Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions

Edited by Jan Borovanský and Patrick A. Riley Melanins and Melanosomes Related Titles Sobin, L. H., Gospodarowicz, M...

Author: Jan Borovanský | Patrick A. Riley | Editors

80 downloads 1228 Views 5MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!

Report copyright / DMCA form

DOWNLOAD PDF

Edited by Jan Borovanský and Patrick A. Riley Melanins and Melanosomes

Related Titles

Sobin, L. H., Gospodarowicz, M. K., Wittekind, C. (eds.)

TNM Classiﬁcation of Malignant Tumours 2009 ISBN: 978-1-4443-3241-4

Rajpar, S., Marsden, J.

ABC of Skin Cancer 2008 ISBN: 978-1-4051-6219-7

Schwartz, R. A.

Skin Cancer 2008 ISBN: 978-1-4051-5961-6

Nordlund, J. J., Boissy, R. E., Hearing, V. J., King, R. A., Oetting, W. S., Ortonne, J.-P. (eds.)

The Pigmentary System Physiology and Pathophysiology 2006 ISBN: 978-1-4051-2034-0

Edited by Jan Borovanský and Patrick A. Riley

Melanins and Melanosomes Biosynthesis, Biogenesis, Physiological, and Pathological Functions

The Editors Prof. Patrick A. Riley Em. Prof. of Cell Pathology University College London Grange Avenue London N20 8AB United Kingdom Prof. Jan Borovanský Charles University 1st Faculty of Medicine Institute of Biochemistry and Experimental Oncology U Nemocnice 5 128 53 Praha 2 Czech Republic Cover The cover design is intended to reﬂect the two principal elements of the book: melanins and melanosomes. The melanins are represented by the graduated background pigment spanning the range of colours from essentially black eumelanin, through reddish to paler shades, representing pheomelanin. The celluar organelles responsible for the production, containment and distribution of melanins, the melanosomes, are represented by symbolic images based on their electron microscopic appearance.

Limit of Liability/Disclaimer of Warranty: While the publisher and authors have used their best efforts in preparing this book, they make no representations or warranties with respect to the accuracy or completeness of the contents of this book and speciﬁcally disclaim any implied warranties of merchantability or ﬁtness for a particular purpose. No warranty can be created or extended by sales representatives or written sales materials. The Advice and strategies contained herein may not be suitable for your situation. You should consult with a professional where appropriate. Neither the publisher nor authors shall be liable for any loss of proﬁt or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. Library of Congress Card No.: applied for British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library. Bibliographic information published by the Deutsche Nationalbibliothek The Deutsche Nationalbibliothek lists this publication in the Deutsche Nationalbibliograﬁe; detailed bibliographic data are available on the Internet at . © 2011 Wiley-VCH Verlag & Co. KGaA, Boschstr. 12, 69469 Weinheim, Germany Wiley-Blackwell is an imprint of John Wiley & Sons, formed by the merger of Wiley’s global Scientiﬁc, Technical, and Medical business with Blackwell Publishing. All rights reserved (including those of translation into other languages). No part of this book may be reproduced in any form – by photoprinting, microﬁlm, or any other means – nor transmitted or translated into a machine language without written permission from the publishers. Registered names, trademarks, etc. used in this book, even when not speciﬁcally marked as such, are not to be considered unprotected by law. Composition Toppan Best-set Premedia Ltd., Hong Kong Printing and Binding Cover Design Graﬁk-Design Schulz, Fußgönheim Printed in the Federal Republic of Germany Printed on acid-free paper ISBN: 978-3-527-32892-5 ISBN oBook: 978-3-527-63615-0 ISBN ePDF: 978-3-527-63614-3

V

Dedication “All nature is but art, unknown to thee; All chance, direction which thou canst not see” Alexander Pope: An Essay on Man Were we to ascribe to chance the existence of this volume we should have to begin with the moment in July 1952 when Professor A.F. Richter, Head of the Second Institute of Medical Chemistry at Charles University in Prague, opened a dust-covered cabinet from which he took, apparently at random, a bottle containing a dark powder and handed it to a young assistant with the words: “Young man, study the contents of this ﬂask.” The assistant was Jiri Duchon and the label on the ﬂask read: “Human melanosarcoma, prepared by H. Waelsch.” Jiri Duchon was born on 27 July 1927, the only son of an eminent scientist. On graduating in medicine in 1952 he joined Richter’s laboratory and his careful analysis of the sample given to him set him upon the course of studies that were to occupy him for the rest of his life. He defended his PhD thesis in 1962 on the topic of “Urinary melanogens in melanoma” and he subsequently made many important contributions to quantitative analysis of the products of melanogenesis. In recognition of his early work, Jiri Duchon was awarded a Roosevelt Fellowship that enabled him to spend 15 months at Harvard in the laboratory of T.B. Fitzpatrick. This was in 1967–1968 when he met and established a friendship with Makoto Seiji who had just developed the methods for melanosome isolation. On his return to Prague, Jiri Duchon set about improving the isolation technique and analyzing these newly discovered organelles. Under his direction and inspiration the Prague laboratory became the leading European center for the detailed biochemical investigation of melanosomes. Jiri Duchon was Head of the Institute for 26 years and many of his collaborators have continued to contribute signiﬁcantly to the ﬁeld of study that he promoted. Professor Duchon was an internationally recognized and highly respected member of the pigment cell fraternity, and was elected an Honorary Member of the European Society for Pigment Cell Research in 1998. It was partly in his honor that the scientiﬁc session on “Melanin and Melanosomes” was arranged at the Federation of European Biochemical Societies (FEBS) Congress in 2009, but

VI

Dedication

tragically he was taken ill on the very day of the Symposium. He was full of encouragement for the project that grew out of the meeting, namely that of publishing a deﬁnitive volume devoted to the subject of his academic endeavors, but died on 2 November 2009, long before it was completed. In recognition of his seminal role in the events that led to the production of this book we dedicate this volume to Jiri Duchon with affectionate remembrance of a ﬁne scientist, an inspirational teacher, a kindly and cultivated companion, and a true friend.

Professor Jiri Duchon MD, PhD, DrSc (1927–2009) (Photograph by K. Meister)

VII

Contents

Dedication V Preface XV List of Contributors 1 1.1 1.2 1.3 1.3.1 1.3.2 1.3.3 1.4 1.5

2 2.1 2.2 2.2.1 2.2.1.1 2.2.1.2 2.2.1.3 2.2.2 2.2.2.1 2.2.2.2 2.2.2.3 2.2.2.4 2.2.2.5 2.3

XIX

History of Melanosome Research 1 Jan Borovanský Introduction 1 Melanosome Research in the Pre-Seiji Era 1 Melanosome Research in the Seiji Era 5 Terminology of Melanosomes 5 Ultrastructural and Histochemical Studies 6 Biochemical Studies 7 Melanosome Research in the Post-Seiji Era 9 Other Historical Aspects 11 Acknowledgments 12 References 13 Classical and Nonclassical Melanocytes in Vertebrates 21 Sophie Colombo, Irina Berlin, Véronique Delmas, and Lionel Larue Deﬁnition of Melanogenic Cells 21 Distribution and Function of Melanogenic Cells 24 Classical Melanocytes 25 Melanocytes in the Epidermis 25 Melanocytes in the Dermis 27 Melanophores in Lower Vertebrates 28 Nonclassical Melanocytes 28 Melanocytes of the Eye 28 Melanocytes of the Inner Ear 31 Melanocytes of the Heart 33 Melanocytes of the Brain and Neuromelanins 36 Melanin in Adipose Tissue 37 Embryonic Development of Melanogenic Cells 37

VIII

Contents

2.3.1 2.3.1.1 2.3.1.2 2.3.2 2.3.2.1 2.3.2.2 2.3.2.3 2.3.2.4 2.4 2.4.1 2.4.2 2.4.2.1 2.4.2.2

3 3.1 3.1.1 3.1.2 3.1.3 3.1.4 3.1.5 3.2 3.2.1 3.2.2 3.2.3 3.2.4 3.2.5 3.2.6 3.3 3.3.1 3.3.1.1 3.3.2 3.3.3 3.3.4 3.3.5

4 4.1 4.2

Classical Melanocytes 38 Early Determined Melanoblasts: The Dorsolateral Pathway 38 Late Determined Melanoblasts: A Common Origin with SCPs and the Dorsoventral Migratory Pathway 40 Nonclassical Melanocytes 41 Melanocytes of the Murine Eye 42 Melanocytes of the Murine Heart 44 Other Nonclassical Murine Melanocytes 45 Other Organisms 45 Transfer of Melanin from Classical and Nonclassical Melanocytes 45 Melanosome Transport 46 Melanosome Transfer 46 Melanosome Transfer from Classical Melanocytes 47 Transfer of Melanin from Nonclassical Melanocytes 51 References 52 Biological Chemistry of o-Quinones 63 Patrick A. Riley, Christopher A. Ramsden, and Edward J. Land General Biological Signiﬁcance of o-Quinones 63 Antibiosis 63 Defensive Secretions 64 Balanid Adhesion 64 Cuticular Hardening in Insects 65 Pigmentation 65 o-Quinone Reactivity 66 Structure and Reactivity 66 Reduction 68 Addition Reactions: Intermolecular addition 71 Polymerization 71 Intramolecular Addition (Cyclization) 72 Addition–Elimination (Substitution) Reactions 73 Role of o-Quinones in Melanogenesis 74 Nonenzymatic Formation of Melanogenic Intermediates 74 Contributions from Pulse Radiolysis to the Chemistry of Eumelanogenesis and Pheomelanogenesis 74 Balance between Eumelanogenesis and Pheomelanogenesis 78 Control of Melanogenesis: Phase I Melanogenesis 78 Tyrosinase Activation 78 Tyrosinase Inactivation 79 References 83 Biosynthesis of Melanins 87 José Carlos García-Borrón and M. Concepción Olivares Sánchez Introduction 87 Raper–Mason Pathway 88

Contents

4.2.1 4.2.2 4.2.3 4.3 4.3.1 4.3.2 4.3.2.1 4.3.2.2 4.3.3 4.3.4 4.3.5 4.4 4.4.1 4.4.2 4.4.2.1 4.4.2.2 4.5

5 5.1 5.1.1 5.1.1.1 5.1.1.2 5.1.1.3 5.1.2 5.1.3 5.2 5.2.1 5.2.1.1 5.2.1.2 5.2.1.3 5.2.2 5.2.2.1 5.2.2.2 5.2.2.3 5.2.2.4 5.2.2.5 5.2.3

Phase I Melanogenesis: The Proximal Raper–Mason Pathway – From L-tyrosine to L-dopachrome 88 Distal Melanogenic Steps: From l-Dopachrome to Eumelanins Biosynthesis of Pheomelanins 91 Structural and Functional Properties of the Melanogenic Enzymes 92 Structure of Tyrosinase and Related Proteins 92 Catalytic Cycle of Tyrosinase 95 Cresolase (Tyrosine Hydroxylase) Reaction Cycle 96 Catecholase (Dopa Oxidase) Reaction Cycle 98 Dct/Tyrp2 98 Tyrp1 100 Other Melanosomal Proteins 101 Regulation of the Melanogenic Pathway 102 Eumelanogenesis versus Pheomelanogenesis: Regulation of the Type of Melanin Pigments 102 Regulation of the Amount of Pigment 104 Regulation of Tyrosinase Levels 104 Control of Tyrosinase-Speciﬁc Activity 106 Conclusions and Perspectives 107 Acknowledgments 109 References 109

90

Inhibitors and Enhancers of Melanogenesis 117 Alain Taïeb, Muriel Cario-André, Stefania Briganti, and Mauro Picardo Introduction 117 Melanin Biochemistry 118 Melanin Biosynthesis 118 Tyrosinase Maturation and Degradation 118 Catalytic Site 119 Paracrine Signaling and Regulation of Epidermal Melanogenesis 119 Methods of Study 120 Depigmenting Agents 121 Agents Acting Prior to Melanin Synthesis 121 Transcriptional Inhibition of Melanogenic Enzymes 121 Post-Translational Modiﬁcation of Melanogenic Enzymes 128 Increased Tyrosinase Ubiquination 129 Agents Acting During Melanin Synthesis 129 Interference with Tyrosinase 129 TRP-2 Modulation 136 Interference with Byproduct Production (Antioxidant and Reducing Agents) 136 Interference with the Melanogenic Pathway 138 Peroxidase Inhibitors 139 Agents Acting After Melanin Synthesis 139

IX

X

Contents

5.2.3.1 5.2.3.2 5.3 5.3.1 5.3.1.1 5.3.1.2 5.3.2 5.3.2.1 5.3.2.2 5.3.2.3 5.3.2.4 5.3.2.5 5.3.2.6 5.3.2.7

Inhibitors of Melanosome Transfer 139 Acceleration of Epidermal Turnover 141 Enhancers of Melanogenesis 143 Activation Through Receptor Mechanisms 144 Melanotropic Peptides 144 Cytokines and Growth Factors 145 Non-Receptor-Mediated Activation 147 Forskolin and cAMP 147 Oligonucleotides and p53 Activation 147 Piperin 147 Lipids (Sphingolipids and Prostaglandins) 147 Phospholipase A2 148 PPAR Activators 148 Psoralens and Photosensitizing Agents 148 References 149

6

Structure of Melanins 167 Shosuke Ito, Kazumasa Wakamatsu, Marco d’Ischia, Alessandra Napolitano, and Alessandro Pezzella Introduction 167 Classiﬁcation and General Properties of Melanins 168 Biosynthetic Studies 169 Early Stages of Melanogenesis 169 Late Stages of Eumelanogenesis 171 Late Stages of Pheomelanogenesis 174 Concept of Mixed Melanogenesis 175 Degradative Studies 176 Eumelanins 176 Pheomelanins 178 Analysis of Eumelanins and Pheomelanins 180 Conclusions 180 References 181

6.1 6.2 6.3 6.3.1 6.3.2 6.3.3 6.3.4 6.4 6.4.1 6.4.2 6.5 6.6

7 7.1 7.2 7.3 7.3.1 7.3.2 7.3.3 7.4 7.4.1 7.4.2 7.4.2.1

Properties and Functions of Ocular Melanins and Melanosomes 187 Małgorzata Rózᠨanowska Introduction 187 Biogenesis of Ocular Melanosomes and Melanogenesis 187 Melanin Content in Pigmented Structures of the Eye 190 Melanin Content in the RPE 190 Melanin Content in the Choroid 193 Melanin Content in the Iris 193 Structure of Ocular Melanosomes 194 Morphology of Ocular Melanosomes 195 Molecular Composition of Ocular Melanosomes 196 Melanosomal Proteins 196

Contents

7.4.2.2 7.5 7.5.1 7.5.2 7.6 7.6.1 7.6.2 7.6.3 7.6.4 7.7 7.7.1 7.7.2 7.7.3 7.8 7.9

8

8.1 8.2 8.3 8.4 8.5 8.6 8.7 8.7.1 8.7.2 8.7.3 8.7.4 8.8

9 9.1 9.2

Melanosomal Lipids 197 Role of Ocular Melanin as a Broadband Optical Filter 198 Role of the Iris as a Filter of Light 198 Role of Melanin in Light Transmission Through the RPE and Choroid 200 Antioxidant Properties of Ocular Melanin 201 Scavenging of Free Radicals 201 Quenching of Electronically Excited States of Photosensitizers and Singlet Oxygen 203 Sequestration of Redox-Active Metal Ions 206 Testing Protective Effects of Ocular Melanin in Cultured Cells 207 Pro-Oxidant Properties of Ocular Melanosomes 209 Generation of ROS and Oxidation of Cellular Reductants 210 Pro-Oxidant Effects of Interactions of Melanosomes with Metal Ions 213 Cytotoxic Properties of Aged RPE Melanin Granules and Their Potential Consequences for Retinal Aging and AMD 214 Other Properties of Ocular Melanosomes and Their Implications 216 Conclusions 217 References 218 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease 225 Kay L. Double, Wakako Maruyama, Makoko Naoi, Manfred Gerlach, and Peter Riederer What are Neuromelanins? 225 Phylogenetic Development of Neuromelanin 227 Development and Metabolism of Neuromelanin 227 Structure of Neuromelanin 231 Biological Role of Neuromelanin in the Human Brain 232 Is Neuromelanin Involved in Neurological Disease? 233 Effects of Neuromelanin In Vitro and In Vivo 235 Mechanisms of Neuromelanin Cytotoxicity 235 Neuromelanin Effects on Mitochondrial Function 237 Neuromelanin Effects on the UPS 239 Comparison of the Cytotoxicity of Neuromelanin with Synthetic DA-M 239 Conclusions 241 Acknowledgments 241 References 241 Biogenesis of Melanosomes 247 Cédric Delevoye, Francesca Giordano, Michael S. Marks, and Graça Raposo Introduction 247 Melanosomes: Intracellular Organelles Specialized in Melanin Synthesis 249

XI

XII

Contents

9.2.1 9.2.2 9.3 9.3.1 9.3.2 9.3.3 9.3.3.1 9.3.3.2 9.3.3.3 9.3.3.4 9.3.4 9.3.5 9.3.5.1 9.3.5.2 9.3.5.3 9.3.5.4 9.4 9.4.1 9.4.2 9.4.2.1 9.4.2.2 9.4.3 9.4.4 9.4.5 9.5

10 10.1 10.2 10.2.1 10.2.2 10.2.3 10.3 10.3.1 10.3.1.1 10.3.2 10.3.2.1 10.3.2.2

Melanosomes Are Unique Organelles That Develop through Different Stages 249 Melanosomal Components 251 Endocytic System and Formation of Melanosomes 254 Organelles of the Endocytic Pathway 254 Melanosomes Are LROs but Are Distinct from Lysosomes 256 Pmel17 and Generation of Early-Stage Melanosomes 259 Pmel17 Structure 259 Pmel17 Forms the Fibrillar Matrix upon Which Melanins Deposit 259 Pmel17 Biosynthesis and Amyloid Formation 260 Functional Importance of Fibrillar Melanosomes 262 OA Type 1 and Melanosome Biogenesis 263 Origin of the Melanosome 263 Early-Stage Melanosomes Originate within the Endocytic Pathway 263 Melanosomes Do Not Originate from the ER 265 Melanosomes Segregate from the Endocytic Pathway beyond Stage I Melanosomes 266 Components of Mature Melanosomes Are Sorted from Distinct Endosomal Intermediates 267 Melanosome Maturation: Cargo Sorting to Mature Melanosomes 269 Griscelli Syndrome and CHS 269 HPS 271 Adaptor Protein (AP) Complexes 271 BLOC Complexes 273 Molecular Motors and the Cytoskeleton 276 SNAREs, Rabs, and Other Regulators 277 Lipids 279 Conclusions 279 Acknowledgments 280 References 281 Transport and Distribution of Melanosomes 295 Mireille Van Gele and Jo Lambert Introduction 295 Model Systems to Study Pigment Transport 296 Melanophores from Fish and Amphibians 296 Mammalian Melanocytes 298 RPE Cells 299 Intracellular Melanosome Transport 299 Microtubule-Based transport 300 Kinesin and Dynein 300 Actin-Based Transport 301 MYO5A 301 RAB27A 302

Contents

10.3.2.3 10.3.2.4 10.3.2.5 10.4 10.5 10.5.1 10.5.1.1 10.5.1.2 10.5.1.3 10.5.2 10.5.2.1 10.5.2.2 10.6 10.7

11 11.1 11.2 11.3 11.4 11.4.1 11.4.2 11.4.3 11.5 11.6 11.7 11.7.1 11.7.2 11.7.3 11.8

12 12.1 12.2

MLPH 303 RAB27A–MLPH–MYO5A Tripartite Protein Complex 304 RAB27A as a New MITF Target Gene 306 Melanosome Motility in RPE: The Rab27a–Myrip–Myo7a Tripartite Complex 307 Melanosome Transfer 309 Modes of Transfer 309 Cytophagocytosis 309 Exocytosis 310 Filopodial-Phagocytosis Model 311 Molecular Players 312 PAR-2 and KGF 312 Adhesion Molecules: Cadherins and Lectins 313 Fate of Melanin in the Keratinocyte 313 Conclusions 315 Acknowledgments 316 References 316 Genetics of Melanosome Structure and Function 323 Vincent J. Hearing Introduction 323 Genes Involved in Melanoblast Development, Migration, and Speciﬁcation 324 Genes Involved in Melanocyte Differentiation, Survival, and Proliferation 325 Genes Involved in Regulating Melanocyte Function 327 Regulation of Constitutive Skin, Hair, and Eye Color 330 Hypopigmentation 332 Hyperpigmentation 332 Genes Involved in the Biogenesis of Melanosomes and Other Lysosome-Related Organelles 333 Genes Involved in Melanin Production 334 Genes Involved in Melanosome Movement, Transfer, and Distribution 336 Movement 336 Transfer 337 Distribution 337 Conclusions 338 References 338 Physiological and Pathological Functions of Melanosomes 343 Jan Borovanský and Patrick A. Riley Tissue Concentration of Melanosomes 343 Melanosome Properties and Functions Are Determined by Their Chemical Composition 344

XIII

XIV

Contents

12.3 12.4 12.4.1 12.4.2 12.4.3 12.4.4 12.5 12.5.1 12.5.2 12.5.3 12.6 12.7 12.7.1 12.7.2 12.7.3 12.8 12.9

13 13.1 13.1.1 13.1.2 13.2 13.3

13.4

13.5 13.6 13.7

Functional Microanatomy of the Melanosome 346 Melanosomes as Centers of Free Radical Activity 350 Free Radical Nature of Melanins 350 Radicals and Reactive Species Associated with Melanogenesis 352 Possible Role of Protein-Bound Dopa 355 Melanosomes as a Therapeutic Target 356 Melanosomes as Energy Transducers 358 Photon/Phonon Conversion 359 Photochemical Reactions 360 Sound/Heat Conversion 360 Melanosomes and Metal Ions 360 Afﬁnity of Melanosomes for Polycyclic and Other Compounds 364 Melanoma Detection and Treatment 365 Participation of Melanosomes in Chemoresistance 366 Long-Term Deposition of Compounds in Melanosomes 367 Exploitation of Melanosomal Proteins and Melanin as Speciﬁc Targets in Melanoma Therapy 368 Conclusions 370 Acknowledgments 370 References 371 Dysplastic Nevi as Precursor Melanoma Lesions 383 Stanislav Pavel, Nico P.M. Smit, and Karel Pizinger Nevi as Risk Factors for Melanoma 383 Development of Melanocytic Nevi 383 Description of Dysplastic Nevi 384 Dysplastic Nevi as Precursor Lesions of Melanoma 384 Cytological Differences between Normal Skin Melanocytes and Dysplastic Nevus Cells: Melanosomal and Mitochondrial Aberrations 386 Metabolic Differences between Normal Skin Melanocytes and Dysplastic Nevus Cells: Preference for Pheomelanogenesis in Dysplastic Nevus Cells 387 Pheomelanogenesis as a Possible Cause of Intracellular Oxidative Imbalance 388 Dysplastic Nevus Cells as Senescent Cells 389 Are Dysplastic Nevus Cells a Class of Cells Exhibiting a Mutator Phenotype? 389 References 391 Index 395

XV

Preface “To that small part of ignorance that we arrange and classify we give the name knowledge” Ambrose Bierce This book is entitled Melanin and Melanosomes, and is about pigment and pigmentation. It is important, however, that we bear in mind that, while the primary function of melanocytes is the production of pigment in melanosomes, these cells have other attributes and perform other signiﬁcant functions. Some of these are well recognized, such as the involvement of the retinal pigment epithelium in photoreceptor physiology (detailed in Chapter 7). Another interesting possibility is that melanogenesis may be the source of some of the substrate for dopamine synthesis [1], and melanocytes may have other important neuroendocrine functions as pro-propriomelanocortin processing cells and a source of prostaglandin D synthase (reviewed by Takeda et al. [2]). Some of these actions may go some way to explaining the remarkable anatomical distribution of melanocytes, often in locations that are not illuminated, such as the leptomeninges. However, this volume is devoted to melanin and melanosomes, and is primarily concerned with vertebrate, especially human, pigmentation. We include melanin that is formed by oxidative processes that are enzymatically catalyzed in specialized cells, both the neural crest-derived dendritic melanocytes (named “classical” melanocytes in Chapter 2) and optic cup-derived retinal pigment epithelial cells (“nonclassical” melanocytes), as well as melanin generated by other oxidative pathways, such as the neuromelanin of the midbrain. The importance of this latter pigment, particularly in relation to Parkinson’s disease, is set out in Chapter 8. The enzymatically generated melanin in vertebrates is synthesized and deposited in specialized intracellular organelles, the melanosomes, and this book concentrates on the many aspects of the formation and functions of these organelles. The melanosome is a highly specialized organelle, the history of which owes much to the early work at Charles University under the aegis of Jiri Duchon (1927–2009), to whom this book is dedicated. Many of the important properties of melanosomes were established in Prague in the early 1970s by a series of investigations on isolated and puriﬁed preparations of this organelle, and investigators at Charles University have continued to

XVI

Preface

contribute signiﬁcantly to the advancement of this ﬁeld, and to follow the developments that have taken place in elucidating the structure of melanosomes and the complex biological roles in which they are implicated. This volume grew from a combination of auspicious factors. In 2009, the 34th Congress of the Federation of European Biochemical Societies (FEBS) was organized in Prague. Naturally, with one of us (J.B.) on the Organizing Committee, one of the scientiﬁc sessions was devoted to melanosomes and, in the wake of the discussions at this meeting, it was felt that there was a signiﬁcant body of new data relating to melanosomes that could usefully be assembled in a volume devoted exclusively to this organelle. It was hoped that such an overview, by integrating the diverse aspects of current knowledge, might help to generate a new understanding of the biological role of melanosomes and stimulate novel research effort in this interesting area of study. We have been fortunate in our publisher, Wiley-VCH, who recognized the timely nature of the proposed volume, and we thank our commissioning editor, Gregor Cicchetti, and his team, especially Anne Chassin du Guerny, for their help and encouragement in bringing the project to fruition. Of course, our main thanks go to our panel of distinguished international contributors who have generously given of their time and expertise in preparing the chapters that we hope form a coherent picture of the up-to-date knowledge in the ﬁeld. Last, but not least, this book celebrates a long and fruitful collaboration between the Editors involving many visits between Charles University and University College London. It is a pleasure to acknowledge the assistance of the British Council in enabling these exchanges. We had hoped initially to have the opportunity to arrange the order of the chapters in the light of their ultimate content so that overlapping areas were most rationally ordered to enable the volume to be read more or less in sequence while allowing the, perforce abundant, cross-references to act as a secondary web in a cohesive network. However, pressure of time prevented us from completing this task and readers may ﬁnd it more convenient to skip between the various contributions according to their interests and predilections. In principle, although the topics are inextricably intertwined, we have elected to place the contributions devoted to melanin – its biosynthesis, chemistry, and properties – at the front of the book, and those dealing with melanosomes – their structure, biogenesis, distribution, and properties – in the following chapters. The topic is put into chronological context by a historical Introduction in Chapter 1, in which Jan Borovanský traces the steps in the discovery of the melanosome, illustrated by portraits of the important investigators that took part in these exciting early studies. As this book is directed largely at aspects of human pigmentation, Chapter 2 consists of a detailed overview by Sophie Colombo, Irina Berlin, Véronique Delmas, and Lionel Larue of the specialized cells in vertebrates in which melanin production in melanosomes takes place. In their contribution a distinction is made between “classical” and “nonclassical” melanocytes. Chapter 3, by Patrick Riley,

Preface

Christopher Ramsden, and Edward Land, emphasizes the central role of the generation and reactivity of o-quinones in melanogenesis, and is followed by Chapter 4 in which the biosynthesis of melanins is reviewed by José Carlos Garcia-Borrón and Conchita Olivares Sánchez. Chapter 5, by Alain Taïeb, Muriel Cario-André, Stefania Briganti, and Mauro Picardo, comprises an analysis of inhibitors and enhancers of melanogenesis. The current understanding of the structure of melanins is then reviewed in Chapter 6 by Shosuke Ito, Kasumasa Wakamatsu, Marco d’Ischia, Alessandra Napolitano, and Alessandro Pezzella, and this is followed in Chapter 7 by a description of the properties and functions of ocular melanins and melanosomes by Małgorzata Różanowska. Chapter 8, by Kay Double, Wakako Maruyama, Makako Naoi, Manfred Gerlach, and Peter Riederer, is devoted to the biological role of neuromelanin in the human brain and its importance in Parkinson’s disease. Chapter 9 consists of a detailed review of the biogenesis of melanosomes by Cédric Delevoye, Francesca Giordano, Michael Marks, and Graça Raposo. This is followed in Chapter 10, by Mireille Van Gele and Jo Lambert, by a description of the transport and distribution of melanosomes. The genetics of melanosome structure and function are skillfully summarized in Chapter 11 by Vincent Hearing. Chapter 12, by Jan Borovanský and Patrick Riley, is devoted to the properties and functions of melanosomes, and, in Chapter 13, the abnormalities of melanosomes and melanogenesis in melanoma precursor lesions are discussed by Stan Pavel, Nico Smit, and Karel Pizinger. We ﬁrmly believe that this compilation of expertise embodies a signiﬁcant work of scholarship, and we sincerely hope that the combined wisdom embraced by this volume conveys both the breadth of detailed and exciting knowledge that currently exists about melanin and melanosomes, and also reveals those shadowed areas of doubt and ignorance that await illumination in the future. March 2011

Patrick A. Riley Jan Borovanský

References 1 Eisenhofer, G., Tian, H., Holmes, C.,

Matsunaga, J., Rofﬂer-Tarlov, S., and Hearing, V.J. (2003) Tyrosinase: a developmentally speciﬁc major determinant of peripheral dopamine. FASEB J., 17, 1248–1255.

2 Takeda, K., Takahashi, N.-H., and

Shibahara, S. (2007) Neuroendocrine functions of melanocytes: beyond the skin-deep melanin maker. Tohoku J. Exp. Med., 211, 201–221.

XVII

wwwwwww

XIX

List of Contributors Irina Berlin Institut Curie Developmental Genetics of Melanocytes, INSERM U1021–CNRS UMR3347 Centre Universitaire 91405 Orsay France

Sophie Colombo Institut Curie Developmental Genetics of Melanocytes, INSERM U1021–CNRS UMR3347 Centre Universitaire 91405 Orsay France

Jan Borovanský Charles University First Faculty of Medicine, Institute of Biochemistry and Experimental Oncology U nemocnice 5 128 53 Prague 2 Czech Republic

Marco d’Ischia University of Naples “Federico II” Department of Organic Chemistry and Biochemistry Via Cinthia 4 80126 Naples Italy

Stefania Briganti San Gallicano Dermatological Institute Laboratory of Cutaneous Physiopathology Elio Chianesi 53 00144 Rome Italy Muriel Cario-André Université de Bordeaux INSERM U1035 146 rue Léo Saignat 33076 Bordeaux France

Cédric Delevoye Institut Curie Centre de Recherche Structure and Membrane Compartments 26 Rue d’Ulm 75248 Paris cedex 05 France Véronique Delmas Institut Curie Developmental Genetics of Melanocytes, INSERM U1021–CNRS UMR3347 Centre Universitaire 91405 Orsay France

XX

List of Contributors

Kay L. Double University of New South Wales Neuroscience Research Australia Barker Street Sydney, NSW 2031 Australia

Jo Lambert Ghent University Hospital Department of Dermatology De Pintelaan 185 9000 Ghent Belgium

José Carlos García-Borrón University of Murcia Department of Biochemistry and Molecular Biology Campus de Espinardo 30100 Murcia Spain

Edward J. Land Keele University School of Physical and Geographical Sciences, Lennard-Jones Laboratories Keele Road Keele ST5 5BG UK

Manfred Gerlach University of Würzburg Clinical Neurobiology, Department of Child and Adolescence Psychiatry, Psychosomatics and Psychotherapy Füchsleinstrasse 15 97080 Würzburg Germany

Lionel Larue Institut Curie Developmental Genetics of Melanocytes, INSERM U1021–CNRS UMR3347 Centre Universitaire, 91405 Orsay France

Francesca Giordano Institut Curie Centre de Recherche Structure and Membrane Compartments 26 Rue d’Ulm 75248 Paris cedex 05 France Vincent J. Hearing National Institutes of Health Laboratory of Cell Biology 37 Convent Drive Bethesda, MD 20892 USA Shosuke Ito Fujita Health University School of Health Sciences Department of Chemistry Toyoake Aichi 470-1192 Japan

Michael S. Marks University of Pennsylvania Departments of Pathology and Laboratory Medicine and Physiology, 513 Stellar-Chance Laboratories 422 Curie Boulevard Philadelphia, PA 19104-6100 USA Wakako Maruyama National Research Center for Geriatrics and Gerontology Department of Cognitive Brain Science Obu Aichi 474-8511 Japan

List of Contributors

Makoko Naoi Gifu International Institute of Biotechnology Department of Neurosciences Kakamigahara Gifu 504-0838 Japan

Mauro Picardo San Gallicano Dermatological Institute Laboratory of Cutaneous Physiopathology Elio Chianesi 53 00144 Rome Italy

Alessandra Napolitano University of Naples “Federico II” Department of Organic Chemistry and Biochemistry Via Cinthia 4 80126 Naples Italy

Karel Pizinger Charles University Department of Dermatology Faculty of Medicine Husova 3 306 05 Pilsen Czech Republic

M. Concepción Olivares Sánchez University of Murcia Department of Biochemistry and Molecular Biology Campus de Espinardo 30100 Murcia Spain

Christopher A. Ramsden Keele University School of Physical and Geographical Sciences, Lennard-Jones Laboratories Keele Road Keele ST5 5BG UK

Stanislav Pavel Leiden University Medical Center Department of Dermatology PO Box 9600 2300 RC Leiden The Netherlands and Charles University Department of Dermatology, Faculty of Medicine Husova 3 306 05 Pilsen Czech Republic

Graça Raposo Institut Curie Centre de Recherche Structure and Membrane Compartments 26 Rue d’Ulm 75248 Paris cedex 05 France

Alessandro Pezzella University of Naples “Federico II” Department of Organic Chemistry and Biochemistry Via Cinthia 4 80126 Naples Italy

Peter Riederer University of Würzburg Clinical Neurochemistry, Department of Psychiatry, Psychosomatics and Psychotherapy, and National Parkinson Foundation Centers of Excellence for Neurodegenerative Diseases Research Füchsleinstrasse 15 97080 Würzburg Germany

XXI

XXII

List of Contributors

Patrick A. Riley Totteridge Institute for Advanced Studies The Grange Grange Avenue London N20 8AB UK Małgorzata Róz·anowska Cardiff University School of Optometry and Vision Science Maindy Road Cardiff CF24 4LU UK Nico P.M. Smit Leiden University Medical Center Central Laboratory for Clinical Chemistry Albinusdreef 2 2333 AC Leiden The Netherlands

Alain Taïeb Hôpital St André Department of Dermatology and Pediatric Dermatology CHU de Bordeaux 1 rue Jean Burguet 33077 Bordeaux France Mireille Van Gele Ghent University Hospital Department of Dermatology De Pintelaan 185 9000 Ghent Belgium Kazumasa Wakamatsu Fujita Health University School of Health Sciences Department of Chemistry Toyoake Aichi 470-1192 Japan

1

1 History of Melanosome Research Jan Borovanský

1.1 Introduction

Melanosomes were ﬁrst proposed as speciﬁc organelles, unique to pigment cells, in a preliminary publication that appeared on 30 July 1960 [1]. An announcement had been made at the 21st Annual Meeting of the Society for Investigative Dermatology, at Miami Beach, Florida, USA on 13 June 1960 [2] and the news, that the chemical composition and enzyme activities in melanosomes and mitochondria are completely different, was considered to be of such signiﬁcance that it appeared in a newspaper report (Figure 1.1). Similar data, with an emphasis on terminology, were published in 1963 [3]. This advance was the result of collaborative work between M. Seiji (1926–1982), at that time working at the Department of Dermatology, Harvard Medical School in Boston under the leadership of T.B. Fitzpatrick (1919–2003) (Figure 1.2), and H. Blaschko and M.S.C. Birbeck, with whom Dr Fitzpatrick established scientiﬁc cooperation during his tenure of a Commonwealth Fellowship at the Department of Biochemistry, Radcliffe Inﬁrmary in Oxford. The history of melanosome research can be formally divided into three parts: (i) the pre-Seiji era (prior to 1960), (i) the Seiji era (1960–1982), and (iii) the postSeiji era (1983–).

1.2 Melanosome Research in the Pre-Seiji Era

The ﬁrst description of mammalian pigment cells was published by Gustav Simon in 1841 [4] who observed round and stellate pigment cells in the hair bulbs of pig embryos. It was preceeded in 1838 by Purkynĕ’s description of pigment in the cells of the substantia nigra, which not only drew attention to pigment granules, but also noted the rise in their numbers with age [5]. We have to admire these early reports because their authors, armed only with primitive light microscopes, were able to ascertain that melanin was not diffusely distributed in the cytoplasm Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

2

1 History of Melanosome Research

Figure 1.1 Announcement of the independent status of melanosome in Medical News on 5

July 1960.

of pigmented cells, but was present in the form of discrete aggregates [5, 6] (Figures 1.3 and 1.4). Deciphering the old literature is problematical as authors often fail to distinguish between melanin (the pigment itself), melanoprotein (the natural melanin– protein complex), and melanin granules (the subcellular organelle). If the method of separation is not adequately described, it is difﬁcult to be certain what material was studied and any conclusions can be misleading [8]. The lack of electron microscopic identiﬁcation of isolated material led to many misinterpretations; for example, the “melanopseudoglobulin” studied by Greenstein et al. [9] was later shown to be melanosomes [10] and Bolt’s “melanoprotein” [11], widely used in biophysical studies, turned out to consist of damaged melanosomes [12]. Mason et al. [10] posed the question of whether melanin granules were particles with a speciﬁc structure or consisted of random aggregates of precipitated metabolic

1.2 Melanosome Research in the Pre-Seiji Era

Figure 1.2

Professor Makoto Seiji (left) and Professor Thomas B. Fitzpatrick (right) in 1972.

Figure 1.3

“Chromatophore” from donkey conjuctiva [7].

products. The introduction of electron microscopy was able to resolve this matter and Laxer et al. [13] were able to discern an inner ultrastructure in isolated melanosomes. The ﬁrst clear pictures were obtained only in 1956 [14]. An avalanche of papers in subsequent years brought with it enormous amounts of information on the ultrastructure of melanosomes and its changes during

3

4

1 History of Melanosome Research

Figure 1.4 Cells of substantia nigra containing neuromelanin [5].

melanosome development (good examples are [15–17]). Other papers (reviewed in [18]) brought together ultrastructural and biochemical data that, in combination, laid the basis for the nomenclature of melanosomal ontogenesis. By comparison with the morphological data, biochemical investigations of melanosomes were more modest, mainly due to the fact that ultrastructural data were derived from studies of intact cells or tissues, whereas biochemical research used samples prepared by relatively harsh preparative procedures. These samples sometimes consisted of melanins, or altered melanosomes, or their fragments, usually without any check of their nature or homogeneity [18]. The aim of researchers in the nineteenth century was not to prepare subcellular particles or native melanoproteins, but to separate the colored pigment (“Farbstoff” = melanin in the terminology of that time). The presence of protein in the isolated material was considered an unwanted contaminant [19]. Probably the ﬁrst mild separation protocol was used by J.J. Berzelius [20]. He investigated pigment (melanosomes?) obtained from eye membranes by water extraction, and noticed its insolubility in acids and limited solubility in alkali. Similar mild extraction procedures were used by Landolt [21] and Mörner [22]. The early isolation procedures were reviewed by Waelsch [23]. He studied “natural melanin” from human melanoma metastases and horse choroids, conﬁrmed the presence of protein attached to pigment, and suggested that melanin could be synthesized from the cyclic amino acids present in the protein moiety; this idea has not been

1.3 Melanosome Research in the Seiji Era

abandoned till now. Herrmann and Boss [24] demonstrated dopa oxidase activity in the fraction of melanin granules from ciliary bodies of cattle eyes, but, as their samples were contaminated with mitochondria, they demonstrated the presence of mitochondrial enzyme markers as well. In 1949, du Buy et al. concluded that melanosomes are modiﬁed mitochondria typical of pigment cells [25]. It is interesting that du Buy [26] and other authors [27] did not abandon the mitochondrial theory of melanosome origin even in 1963 (i.e., 2 years after the formulation of Seiji’s melanosomal concept) and even published their papers in the same volume in which Seiji et al. published detailed conﬁrmation of their model [28]. It is interesting that history has disregarded the contribution of Stein [29] who, several years before the work of Seiji et al., using a separation procedure of his own, isolated melanin granules from ox choroids and analyzed their content not only of melanin, but also lipids, carbohydrates, RNA, and metals (including the pioneer ﬁnding of a high level of zinc), and concluded that the chemical composition of melanin granules is completely different from mitochondria. The ability of melanin in melanin granules, isolated from Harding-Passey melanoma and from the ink sac of Loligo opalescens, to act as a cation exchanger [30], and the demonstration of free radical activity in melanin-containing tissues [31] also rank among the observations of the pre-Seiji era.

1.3 Melanosome Research in the Seiji Era 1.3.1 Terminology of Melanosomes

The demonstration of melanosomes as unique pigment cell organelles possessing developmental stages prompted the introduction of a system of terminology that reﬂected the characteristics of the various states. Until 1961 the common term for all varieties of these organelles was melanin (or pigment) granule [1, 2]. The ﬁrst system of nomenclature [2] described three stages in the ontogenesis of melanosomes: i) Premelanosomes: spherical organelles. ii) Melanosomes: organelles with an internal structure and tyrosinase activity. iii) Melanin granules: melanoprotein polymer. A second terminological system was proposed [3, 26] consisting of three developmental stages plus a ﬁnal product. Thus:

• • • •

Stage I (ﬁrst stage): biosynthesis of protein. Stage II (intermediate stage): biosynthesis of organelle. Stage III (late phase): biosynthesis of melanin. Final product: melanin granule.

5

6

1 History of Melanosome Research

These nomenclature systems introduced a certain degree of confusion, particularly as the term melanin granule had been used to describe pigment granules at any developmental stage. In an attempt to establish a consensus, Fitzpatrick et al. [32, 33] circulated a postal questionnaire seeking opinions about the adequacy of the terms in common use in pigment cell research and, with the approval of the participants of the Sixth International Pigment Cell Conference in 1965 in Soﬁa, Bulgaria, recommended the use of two terms:

•

Melanosome: a discrete melanin-containing organelle in which melanization is complete as indicated by its almost uniform density by electron microscopy and the absence of demonstrable tyrosinase activity.

•

Premelanosome: a term applied to all the stages in melanosome biogenesis that precede the fully developed state. Within the restrictions of this general deﬁnition, the premelanosomal stage might, at the discretion of the investigator, be subdivided into early, intermediate, and late phases.

The nomenclature in general use today does not adhere to any of the three systems outlined above, but is essentially a system proposed by Toda et al. [34–36] reﬂecting the earlier descriptions of Birbeck [37, 38] which employs the uniform term “melanosome” with a numerical indication (I–IV) of the degree its ontogenetic development. However, in practice, chaos prevails. While the system of Toda et al. is widely – if somewhat erratically – used, some European authors refer, often incorrectly, to the stages proposed in the second system of nomenclature [3, 26] and some American authors tend to cite nomenclature introduced in their previous papers or those of their friends. 1.3.2 Ultrastructural and Histochemical Studies

The concept of subcellular biosynthesis and localization of melanins and melanoproteins in melanosomes was further conﬁrmed by (i) autoradiographic evidence with [3H]dopa and [2-14C]dopa [39–43]., (ii) incorporation of [2-14C]dopa and monitoring radioactivity in subcellular fractions [44, 45]., and (c) isolation of melanosomes and analysis of their chemical composition [46, 47]. Electron microscopy enabled the deﬁnition of the basic morphometric data of isolated melanosomes (i.e., their size, shape, and ultrastructural appearance). The most extensive data were published by Hach et al. [48, 49]. For discussion concerning the ultrastructural appearances of melanosomes, see Section 12.3 in Chapter 12. Various pathological states may be manifested by changes in melanosome morphology. Mishima et al. [50] considered that melanosome polymorphism, such as changes in size, shape, ultrastructural matrix, the manner of melanin deposition, and the degree of melanosome maturation, as a criterion of molecular pathology that could ﬁnd practical use in the differential diagnosis of various pigmentary disorders.

1.3 Melanosome Research in the Seiji Era

In melanoma cells, various irregularities in the architecture of melanosomes are common. Deposition of melanin can be uneven, leading to a bizarre appearance of melanosomes [51–57]; the presence of melanosomes of all stages of development is typical [51, 52]. Melanoma melanosomes also often exhibit defects of their limiting membranes that may lead to leakage of toxic melanin precursors into the cytosol. The pathological consequences of this failure of containment of melanogenic intermediates are discussed in Section 12.4.2. Extracellular deposition of melanin on ﬁbrils resembling melanosomal matrix ﬁbrils has also been observed in melanoma cells [58, 59]. Early ideas on melanosomal biogenesis were summarized in several studies [52, 60, 61]. Fitzpatrick and Breathnach deﬁned a functional unit in human epidermis named the “epidermal melanin unit.” This was viewed as a symbiotic relationship between melanocytes and keratinocytes in which each melanocyte supplies approximately 36 keratinocytes with melanosomes [62]. The mechanism of melanosome transport and transfer to keratinocytes was outlined in Mottaz and Zelickson [63]. Szabó et al. [64] demonstrated racial differences in the fate of melanosomes in human epidermis. 1.3.3 Biochemical Studies

A prerequisite for classical biochemical studies is the ability to isolate native and pure melanosomes [65]. To this end 17 isolation protocols suggested for the isolation of melanosomes between 1940 and 1973 were critically reproduced [18, 65], and it was concluded that the best samples could be obtained using the procedures described by Stein [29], Doezema [66], and Haberman and Menon [67]. Isolation of melanosomes from keratinous material turned out to be more difﬁcult because the need to release melanosomes from hair required chemical means of tissue disintegration, which always engenders a search for a compromise between sufﬁcient tissue disintegration and minimizing the extent of melanosome modiﬁcation by the isolation procedure [61, 68]. Ten methods for isolating melanosomes from hair were critically reviewed and half of them, which from the assessment of the isolation conditions seemed to be promising, were reproduced [68]. The best results were obtained with the isolation protocol of Borovanský and Hach [69]. The availability of melanosome samples of adequate quality [18] opened the gate to the subsequent establishment of their basic chemical composition. Melanin and protein moieties were shown to be dominant constituents of melanosomes [46, 47, 61, 70–72]. Isolated melanosomes were reported to contain 5–10% of carbohydrates [29]. Tyrosinase, as a glycoprotein, also brings into melanosomes sialic acids containing N-acetylneuraminic acid [73]. In addition to gangliosides mentioned in Section 12.2, many other lipid constituents in melanosomes have been reported including cholesterol and free fatty acids [74], and phospholipids [3, 75]. The level of total lipids was found to vary between 1–5% [29] and 5–11% [76]. Jimbow et al. [77] made a complete qualitative and quantitative analysis of lipids and their

7

8

1 History of Melanosome Research

Figure 1.5 Professor Kowichi Jimbow, a pioneer in melanosome research, in 1981.

fractions in isolated Harding-Passey and B16 melanoma melanosomes, and found quantitative differences between them (Figure 1.5). The demonstration of the absence of DNA in isolated melanosomes was clear evidence of the difference between melanosomes and mitochondria [78]. Melanosomes are abundant in various metals as described in detail in the Section 12.6. The description of the development of analytical methods including classical chemical techniques such as titration, spectrophotometry, electrochemical and isotope methods, neutron activation analysis, mass spectrometry, and inductively coupled plasma techniques, together with cell biological methods such as histochemistry, autometallography, autoradiography, and microanalytical techniques (using electron, proton, laser, X-ray, and ion beams), and their use in melanosome analyses is comprehensively covered in a review [79]. Nowadays it sounds incredible, but in the 1960s there were doubts as to whether melanosomes merely represented an association of tyrosinase with melanin or whether the organelles contained other proteins as suggested by the electron microscopic appearance. The matter was investigated by electrophoretic studies of isolated melanosomes treated with detergents. Of the many studies summarized in [61], only four used samples of melanosomes that had been checked for purity by electron microscopy [66, 80–82] and these publications unambiguously demonstrated that melanosomes contain several proteins, some of them of brown color and rapid anionic mobility on electrophoresis suggesting their melanoprotein nature [80–82]. The presence of more proteins in melanosomes was later conﬁrmed by comparison of the amino acid composition of tyrosinase with that of melanosome hydrolysates [11]. Matrix proteins were characterized by means of sodium dodecyl sulfate–polyacrylamide gel electrophoresis in melanosomes isolated from Harding-Passey and B16 melanomas and treated with Brij-35 and

1.4 Melanosome Research in the Post-Seiji Era

guanidine hydrochloride. A simultaneous ultrastructural study revealed that treatment of melanosomes with guanidine hydrochloride induced partial degradation detectable by electron microscopy [83]. A strong stream of research represented studies aimed at demonstrating the presence of enzymes in melanosomes. Naturally, special attention was paid to the melanosomal marker enzyme – tyrosinase (EC1.14.18.1) [1–3, 84–89]. Among the common constituents of melanosomes are acid phosphatase (EC 3.1.3.2.) [90–95] and other lysosomal hydrolases, such as β-galactosidase (EC 3.2.1.23) [75], β-glucuronidase (EC 3.2.1.31) [74, 96], β-N-acetyl glucosaminidase (EC 3.2.1.30) [74], cathepsin D (EC 3.4.23.5) [74], and arylsulfatase (EC 3.1.6.1) [97]. The presence of acid phosphatase and other acid hydrolases used to be explained by adhesion of lysosomal enzymes because during isolation melanosomes are contaminated with phagosomes [90, 96] and autophagosomes [94, 97]. However, as removal of superﬁcially bound proteins by detergents [96] or enzymes [74] did not remove the activity of lysosomal enzymes, they seemed to be integral constituents of melanosomes. Tyrosine-2-oxoglutarate amino transferase (EC 2.6.1.5) and tryptophan-2,3dioxygenase (EC1.13.11.11) were demonstrated to be a constant constituent of melanosomes from guinea pig skin [98]. The presence of ATPase (EC 3.6.1.3) is not surprising [45]. γ-Glutamyltransferase (EC 2.3.2.2) was demonstrated in melanosomes and premelanosomes of B16 melanoma cells [99]. γ-Glutamyltransferase is thought to have a role both in melanogenesis and in cellular protection against oxidative stress. Progress in melanosome research was quite rapid. Ten years after the recognition of the melanosome as a unique subcellular particle of pigment cells, the basic biological processes associated with pigmentation were shown to be related to: (i) formation of melanosomes in melanocytes, (ii) melanization of melanosomes in melanocytes, (iii) transfer of melanosomes into keratinocytes, and (iv) transport of melanosomes by keratinocytes, with or without degradation, in lysosome-like organelles [100]. These four processes were partially characterized, and biochemical knowledge of melanosomes reached a level enabling consideration of their function and the possibilities of exploiting these functions in clinical practice [101–105]. A well-balanced review on the melanosome and melanogenesis, describing the situation at the beginning of the twenty-ﬁrst century, was written by Tolesson [106]. The advent of the techniques of molecular biology has still further accelerated the growth of our knowledge of melanosomes.

1.4 Melanosome Research in the Post-Seiji Era

Professor Makoto Seiji died in 1982. In recognition of his key role and fundamental achievements in melanosome research the Seiji Memorial Lectureship was established in his memory by the International Federation of Pigment Cell Societies to be given every third year at the International Pigment Cell Conferences.

9

10

1 History of Melanosome Research

Symbolically, the ﬁrst Seiji Memorial Lecture was given by Professor T.B. Fitzpatrick at the International Pigment Cell Conference in Giessen in 1983. In parallel with the “epidermal melanin unit” a “follicular melanin unit” was introduced [107]. Modern analytical techniques of high sensitivity make heavy demands on the purity of the melanosomal fractions studied. Hence, the problem of isolation has resurfaced. For the isolation of melanosomes from keratinized structures enzymatic tissue disintegration has been introduced by Arnaud and Boré [108]. However, they used preliminary treatment of hair either with dimethylsulfoxide at 120 °C or treatment of hair under reﬂux with an aqueous solution of lithium bromide. Such methods are in absolute contradiction to principles of denaturationfree separation. Isolation methods strongly predetermine the quality of the samples obtained, such as surface area-to-mass ratio as demonstrated by Liu and Simon [109]. In 2000, Prota et al. developed an isolation procedure based only on enzyme digestion [110]. Melanosomes of various stages could be separately isolated by inserting into the protocol a free-ﬂow electrophoresis step [111]. Percoll gradients were also introduced into melanosome isolation [112]. Tyrosine-induced increase of melanin was shown to inﬂuence melanosomal size and shape, especially of those originating from the light skin types [113]. The list of lysosomal hydrolases was extended by the detection of cathepsin B (EC 3.4.22.1) and L (EC 3.4.22.15) [114], and α-mannosidase (EC 3.2.1.24) [99]. After the discovery of the acidic pH of melanosomes [115], the presence both of acid hydrolases and lysosome-associated membrane proteins 1, 2, and 3, and evidence of phagocytotic ability, melanosomes were designated as specialized lysosomes [116, 117] and their existence as speciﬁc organelles was endangered. The ranking of melanosomes among lysosome-related organelles [118] was the only appropriate solution, which readily explains the common participation of lysosomes and melanosomes in some pigmentary disorders such as Chediak–Higashi syndrome and Heřmanský–Pudlák syndrome. The mechanism of melanosome disintegration and degradation has been studied for a long time [119]. There are many histochemical reports of the presence of acid hydrolases, and particularly acid phosphatase, in melanosome complexes and these have been interpreted as implying their presumptive role in melanosome degradation (reviewed in [119, 120]). However, the reaction speciﬁcity of acid phosphatase consists of hydrolyzing phosphate esters and there have been no reports to suggest that phosphates play any part in maintaining the aggregation pattern of melanin or melanosome architecture. Acid hydrolases may have a role the degradation of the protein moiety of the melanosome, but melanin seems to be susceptible mainly to redox reactions [119–122] (see also Sections 12.4.1 and 12.4.2). Immunological techniques have helped a great deal in understanding melanosome structure and biogenesis. Monoclonal antibodies prepared by immunization with melanosomal proteins [123], but especially antibodies prepared by Hearing against synthetic peptides corresponding to the C-termini of melanosomal proteins, have proved to be invaluable tools in melanosome research [124]. In the post-Seiji era the contribution of molecular biological techniques has been enormous and is reﬂected in Chapters 2, 9, 10, and 11. The group of Professor John

1.5 Other Historical Aspects

Simon has recently introduced new sophisticated biophysical and chemical techniques into melanosome research (e.g., [109, 125]). The combined consensus of the current knowledge of melanins and melanosomes that has emerged from the many investigations brieﬂy alluded to above constitutes the material contained within the chapters written by the leading authorities in the ﬁeld that illuminate this book.

1.5 Other Historical Aspects

The author has been engaged in pigment research since 1968 and this chapter reﬂects his subjective preferences for the articles taking into account melanosomes as subcellular organelles. Hundreds of articles (and their authors) dealing with the investigation of processes, control factors, and molecular characteristics of melanocytes, which have no direct relation to melanosomes as functional units, can be found in other reviews. The description of pigment cell research along a time axis was monitored in a unique way by Nordlund et al. [126] and there are also articles with a geographical emphasis on pigment cell research [127, 128]. The history of melanocyte research, mentioning the ﬁrst description by Sangiovanni in 1819 [129] of a pigment cell as a “chromatophore” in the squid, was summarized by Westerhof [130] and repeated by Falabella [131]. Brief historical remarks can be found in [132, 133]. Melanoproteins were ﬁrst deﬁned in 1910 [134], and studied again by Serra [135] and reviewed in [6, 136]. Since the formulation of an exact deﬁnition of speciﬁc melanosomal proteins the general term “melanoprotein” has been fading. However, the terminology reappears on occasion in descriptions of the manner of melanin attachment to proteins such as Pmel-17. Of course, the history of melanin and the development of knowledge in the ﬁeld is much longer than the time since it was given its name by Berzelius in 1840 [20], and is covered in considerable depth in the books by Nicolaus [137] and Prota [138], and in several reviews (e.g., [139]). In his book, Nicolaus [137] divides the development of melanin chemistry into three periods. (i) The period of frustration, which started with the studies of Dressler and Pribram in 1856 and terminated with Raper’s fundamental work in the 1930s. (ii) The period of uncertainty 1930–?. In 1968, Nicolaus predicted that ever-increasing interest would soon lead to entry into the third period – (iii) the period of elucidation. It is undoubtedly to this era that the articles of this book belong. The ever-increasing interest in the investigation of melanosomes can be illustrated by data from the ISI Web of Knowledge (Table 1.1). Until 1960 the term melanosome on the ISI Web of Knowledge did not exist and a slow increase took place up to the period 1981–1985. The decrease in the subsequent period can be explained by the failure to use the term melanosome among the “key words” as investigators concentrated more on the molecular level. A further complication is that the term melanosome has two meanings: (i) a subcellular particle of pigment cells as described in this book and (ii) a dark region

11

12

1 History of Melanosome Research

Figure 1.6 Melanosome – a dark region present in migmatite rocks – on the staircase of the Institute of Biochemistry and Experimental Oncology, First Faculty of Medicine, Charles University in Prague. Length = 54 mm. Table 1.1 Number of entries under the term “melanosome” in the ISI Web of Knowledge (www.isiknowledge.com).

Period (years)

No. entries

1961–1965 1966–1970 1971–1975 1976–1980 1981–1985 1986–1990 1991–1995 1996–2000 2001–2005 2006–2010

4 13 26 29 49 33 131 167 289 343

present in migmatite rocks [140] (Figure 1.6). The entries in Table 1.1 have been adjusted to exclude the geological citations.

Acknowledgments

Supported by VZ MSTM CR 0021620808 and IGA MZ NT11229-3. Thanks belong to Professor Yasushi Tomita from Nagoya University in Japan, who kindly supplied two photographs (Figures 1.1 and 1.2) from his private archive.

References

References 1 Baker, R.V., Birbeck, M.S., Blaschko, H.,

2

3

4

5

6

7

8

9

10

11

Fitzpatrick, B., and Seiji, M. (1960) Melanin granules and mitochondria. Nature, 187, 392–394. Seiji, M., Fitzpatrick, T.B., and Birbeck, M.S.C. (1961) The melanosome: a distinctive subcellular particle of mammalian melanocytes and the site of melanogenesis. J. Invest. Dermatol., 36, 243–252. Seji, M., Fitzpatrick, T.B., Simpson, R.T., and Birbeck, M.S.C. (1963) Chemical composition and terminology of specialized organelles (melanosomes and melanin granules) in mammalian melanocytes. Nature, 197, 1082–1084. Simon, G. (1841) Zür Entwickelungsgeschichte der Haare. Joh. Muller’s Arch. Anat., 367. Purkynĕ, J.E. (1838) Bericht über die Versammlung deutscher Naturforscher und Aerzte in Prag im September 1837 (eds K. Sternberg and J.V. Krombholz), Prague, pp. 174–180. Sorby, H.C. (1878) On the colouring matters found in human hair. J. Anthropol. Inst., 8, 1–24. Kromayer, E. (1893) Oberhautpigment der Säugethiere. Arch. Mikrosk. Anat., 42, 1–15. Duchoň, J., Fitzpatrick, T.B., and Seiji, M. (1968) Melanin 1968: some deﬁnitions and problems, in The 1967–68 Year Book of Dermatology (eds A.W. Kopf and R. Andrade), Year Book Medical, Chicago, IL, pp. 6–33. Greenstein, J.P., Turner, J.C., and Jenrette, W.V. (1940) Chemical studies on the components of normal and neoplastic tissues. IV. The melanincontaining melanopseudoglobulin of the malignant melanoma of mice. J. Nat. Cancer Inst., 1, 377–385. Mason, H.S., Kahler, E., Mac Cardle, R.C., and Dalton, A.J. (1947) Chemistry of melanin. IV. Electron micrography of natural melanins. Proc. Soc. Exp. Biol. Med., 66, 421–431. Bolt, A.G. (1967) Interactions between human melanoprotein and chlorpromazine derivatives. I. Isolation

12

13

14

15

16

17

18

19

20

21

22

and puriﬁcation of human melanoprotein from hair and melanoma tissue. Life Sci., 6, 1277–1283. Borovanský, J. and Hach, P. (1973) Comments on Bolt’s tumour melanoprotein. Neoplasma, 20, 325–329. Laxer, G., Sikorski, J., Whewell, C.S., and Woods, H.J. (1954) The electron microscopy of melanin granules isolated from pigmented mammalian ﬁbres. Biochim. Biophys. Acta, 15, 174–185. Birbeck, M.S.C., Mercer, E.H., and Barnicot, N.A. (1956) The structure and formation of pigment granules in human hair. Exp. Cell Res., 10, 505–514. Wellings, S.R. and Siegel, J. (1959) Role of Golgi apparatus in the formation of melanin granules in human malignant melanoma. J. Natl. Cancer Inst., 3, 131–140. Drochmans, P. (1960) Electron microscope studies of epidermal melanocytes, and the ﬁne structure of melanin granules. J. Biophys. Biochem. Cytol., 8, 165–180. Drochmans, P. (1960) Study by the electron microscope of the mechanism of melanin pigmentation. Arch. Belg. Dermatol. Syphiligr., 16, 155–163. Borovanský, J. (1975) Isolation of melanosomes and an attempt to quantify melanin content in tissues. PhD thesis, Faculty of General Medicine, Charles University Prague. Sieber, N. (1886) Ueber die Pigmente der Chorioidea und der Haare. Arch. f. Exp. Pathol., 20, 362–367. Berzelius, J.J. (1840) Lehrbuch der Chemie (aus der Schwedischen Handschrift des Verfassers übersetzt von F. Woehler). Dritte ungearbeitete und vermehrte Original Auﬂage, Dresden & Leipzig: in der Arnoldischen Buchhandlung, vol. 9, 22–24. Landolt, H. (1899) Ueber das Melanin der Augenhäute. Hoppe Seylers Z. Physiol. Chem., 28, 192–211. Mörner, K.A.H. (1887) Zur Kenntnis von der Farbstoffen der melanotischen Geschwülste. Z. Physiol. Chem., 11, 66–141.

13

14

1 History of Melanosome Research 23 Waelsch, H. (1932) Zur Kenntnis der

24

25

26

27

28

29

30

31

32

33

natürlichen Melanine. Hoppe Seylers Z. Physiol. Chem., 213, 35–57. Hermann, H. and Boss, M.B. (1945) Dopa oxidase activity in extracts from ciliary body and in isolated pigment granules. J. Cell. Comp. Physiol., 26, 131–138. Du Buy, H.G., Woods, M.W., Burk, D., and Lackey, M.D. (1949) Enzymatic activities of isolated amelanotic and melanotic granules of mouse melanomas and suggested relationship with mitochondria. J. Am. Cancer Inst., 9, 325–336. Du Buy, H.G., Showacre, J.L., and Hesselbach, M.L. (1963) Enzymic and other similarities of melanoma granules and mitochondria. Ann. NY Acad. Sci., 100, 569–583. Woods, M., Burk, D., and Hunter, J. (1963) The ontogenic status of melanin granules. Ann. NY Acad. Sci., 100, 534–539. Seiji, M., Shimao, K., Birbeck, M.S.C., and Fitzpatrick, T.B. (1963) Subcellular localization of melanin biosynthesis. Ann. NY Acad. Sci., 100, 497–533. Stein, W.D. (1955) Chemical composition of the melanin granule and its relation to the mitochondrion. Nature, 175, 256–257. White, L.P. (1956) Melanin: a naturally occurring cation exchange material. Nature, 182, 1427–1428. Commoner, J.B., Townsend, J., and Pake, G.E. (1954) Free radicals in biological materials. Nature, 174, 689–691. Fitzpatrick, T.B., Quevedo, W.C., Jr, Levene, A.L., Mc Govern, V.J., Mishima, Y., and Oettle, A.G. (1966) Terminology of vertebrate melanin-containing cells. Science, 152, 88–89. Fitzpatrick, T.B., Quevedo, W.C., Levene, A.L., Mc Govern, V.J., Mishima, Y., and Oettle, A.G. (1966) Terminology of vertebrate melanin-containing cells, their precursors and related cells. A report of the nomenclature committee of the 6th International Pigment Cell Conference, in Structure and Control of the Melanocyte (eds G. Della Porta and O. Muhlbock), Springer, New York, pp. 1–5.

34 Toda, K., Hori, Y., and Fitzpatrick, T.B.

35

36

37

38

39

40

41

42

43

(1968) Isolation of the intermediate “vesicles” during ontogeny of melanosomes in embryonic chick retinal pigment epithelium. Fed. Proc., 27, 722. Toda, K. and Fitzpatrick, T.B. (1971) The origin of melanosomes, in Biology of Normal and Abnormal Melanocytes (eds T. Kawamura, T.B. Fitpatrick, and M. Seiji), University Park Press, Baltimore, MD, pp. 265–278. Fitzpatrick, T.B., Hori, Y., Toda, K., Kinebuchi, S., and Szabó, G. (1971) The mechanism of normal human melanin pigmentation and of some pigmentary disorders, in Biology of Normal and Abnormal Melanocytes (eds T. Kawamura, T.B. Fitpatrick, and M. Seiji), University Park Press, Baltimore, MD, pp. 369–401. Birbeck, M.S.C. (1962) Electron microscopy of melanocytes. Br. Med. Bull., 18, 220–222. Birbeck, M.S.C. (1963) Electron microscopy of melanocytes: the ﬁne structure of hair bulb premelanosomes. Ann. NY Acad. Sci., 100, 540–548. Brumbaugh, J.A. and Froiland, T.G. (1973) DOPA and cysteine incorporation into premelanosomes: effects of cycloheximide and gene substitution. J. Invest. Dermatol., 60, 172–178. Hempel, K. and Deimel, M. (1963) Untersuchungen zur gezielten Strahlentherapie des Melanoms und des chromafﬁnen Systems durch selektive 3 H-Inkorporation nach Gabe von 3H markierten DOPA. Strahlentherapie, 121, 22–45. Hempel, K. (1966) Investigation on the structure of melanin in malignant melanoma with 3H and C14-DOPA labeled at different positions, in Structure and Control of the Melanocyte (eds G. Della Porta and O. Muhlbock), Springer, New York, pp. 162–175. Nakai, T. and Shubik, P. (1964) Electronmicroscopic radioautoraphy: the melanosome as a site of melanogenesis in neoplastic melanocytes. J. Invest. Dermatol., 43, 267–269. Zelickson, A.S., Hirsch, H.M., and Hartmann, J.F. (1964) Melanogenesis – an autoradiographic

References

44

45

46

47

48

49

50

51

52

53

54

55

study at the ultrastructural level. J. Invest. Dermatol., 43, 327- 332. Seiji, M. and Iwashita, S. (1963) On the site of melanin formation in melanocytes. J. Biochem., 54, 465–467. Seiji, M. and Iwashita, S. (1965) Intracellular organisation of tyrosinase and site of melanin formation in melanocyte. J. Invest. Dermatol., 45, 305–314. Borovanský, J. and Duchoň, J. (1974) Chemical composition of hair melanosomes. Dermatologica, 149, 116–120. Hach, P., Borovanský, J. and Duchoň, J. (1973) Melanosomes of horse benign melanoma. Folia Morphol. (Prague), 21, 275–277. Hach, P. and Borovanský, J. (1972) Ultrastructure of melanosomes of different origin. Folia Morphol. (Prague), 20, 82–84. Hach, P., Duchoň, J., and Borovanský, J. (1977) Ultrastructural and biochemical characteristics of isolated melanosomes. Folia Morphol. (Prague), 25, 407–410. Mishima, Y. (1965) Macromolecular changes in pigmentary disorders. Arch. Dermatol., 91, 519–557. Césarini, J.P. (1971) Recent advances in the ultrastructure of malignant melanoma. Rev. Eur. Étud. Clin. Biol., 16, 316–322. Foa, C. and Aubert, C.H. (1977) Cellular localization of tyrosinase in human malignant melanoma. J. Invest. Dermatol., 68, 369–378. Hirone, T., Nagai, T., Matsubara, T., and Fukushiro, R. (1971) Human malignant melanoma of the skin and their preexisting conditions, in Biology of Normal and Abnormal Melanocytes (eds T. Kawamura, T.B. Fitpatrick, and M. Seiji), University Park Press, Baltimore, MD, pp. 329–348. Hunter, J.A.A., Paterson, W.D., and Fairley, D.J. (1978) Human malignant melanoma. Melanosomal polymorphism and the ultrastructural DOPA reaction. Br. J. Dermatol., 98, 381–390. Hunter, J.A.A., Zaynoun, W.D., Paterson, W.D., Bleehen, S.S., Mackie, R., and Cochran, A.J. (1978) Cellular ﬁne structure in the invasive nodules of

56

57

58

59

60

61

62

63

64

65

66

67

different histogenetic types of malignant melanoma. Br. J. Dermatol., 98, 255–272. Moyer, F.H. (1963) Genetic effects on melanosome ﬁne structure and ontogeny in normal and malignant cells. Ann. NY Acad. Sci., 100, 584–606. Szekeres, L. (1975) Fine structure and X-ray microanalysis of melanosomes in pigmented nevi and melanoma. Arch. Derm. Forsch., 252, 297–304. Klingmuller, G. and Schmoeckel, C. (1971) Frei im Cytoplasma liegende Melanin-synthesierende Membranaordnungen beim malignem Melanom. Arch. Derm. Forsch., 241, 115–121. McGovern, V.J. and Lane Brown, M.M. (1969) The Nature of Melanoma, Thomas, Springﬁeld, IL. Fitzpatrick, T.B. (1971) The biology of pigmentation. Birth Defects Orig. Artic. Ser., 7, 5–12. Borovanský, J. (1978) Biochemical parameters of melanosomes and pigment tissues. Habilitation thesis. Charles University, Prague. Fitzpatrick, T.B. and Breathnach, A.S. (1963) Das epidermale melanin einheit-system. Dermatol. Wochenschr., 147, 481–489. Mottaz, J.H. and Zelickson, A.S. (1967) Melanin transfer: a possible phagocytic process. J. Invest. Dermatol., 49, 605–610. Szabó, G., Gerald, A.B., Pathak, M.A., and Fitzpatrick, T.B. (1969) Racial differences in the fate of melanosomes in human epidermis. Nature, 222, 1081–1082. Borovanský, J., Hach, P., Vedralová, E., and Duchoň, J. (1978) Strategy of melanosome isolation, in XXIst Colloqiuum Scientiﬁcum Facultatis Medicae Universitatis Carolinae et XIXth Morphological Congress Symposia (ed. E. Klika), Univerzita Karlova, Prague, pp. 613–618. Doezema, P. (1973) Proteins from melanosomes of mouse and chick pigment cells. J. Cell. Physiol., 89, 201–208. Habermann, H.F. and Menon, I.A. (1973) A modiﬁed method for the isolation of melanosomes from B-16

15

16

1 History of Melanosome Research

68

69

70

71

72

73

74

75

76

77

melanoma. J. Invest. Dermatol., 60, 67–72. Borovanský, J. and Hach, P. (1986) Isolation of melanosomes from keratinous structures: current state of the art. Arch. Dermatol. Res., 279, 54–58. Borovanský, J. and Hach, P. (1972) Isolation of melanosomes from keratinous material – a new method. Dermatologica, 145, 37–41. Duchoň, J., Borovanský, J., and Hach, P. (1973) Chemical composition of ten kinds of various melanosomes, in Mechanisms in Pigmentation (eds V.J. McGovern and P. Russell), Karger, Basel, pp. 165–170. Ito, S. and Jimbow, K. (1983) Quantitative analysis of eumelanin and pheomelanin in hair and melanomas. J. Invest. Dermatol., 80, 268–272. Jimbow, K., Miyake, Y., Homma, K., Yasuda, K., Izumi, Y., Tsutsumi, A., and Ito, S. (1984) Characterization of melanogenesis and morphogenesis of melanosomes by physicochemical properties of melanin and melanosomes in malignant melanoma. Cancer Res., 44, 1128–1134. Miyazaki, K. and Ohtaki, N. (1975) Tyrosinase as a glycoprotein. Arch. Dermatol. Forsch., 252, 211–216. Siakotos, A.N., Patel, V., and Cantaboni, A. (1973) The isolation and chemical composition of premelanosomes and melanosomes: human and mouse melanomas. Biochem. Med., 7, 14–24. Seiji, M. (1966) Subcellular particles and melanin formation in melanocytes, in Advances in the Biology of Skin VIII: The Pigmentary System (eds W. Montagna and F. Hu), Pergamon Press, Oxford, pp. 189–222. Vedralová, E. and Duchoň, J. (1977) Lipid constituents in melanosomes of tumour origin. Sborník lék., 79, 335–339. Jimbow, M., Kanoh, H., and Jimbow, K. (1982) Characterization of biochemical properties of melanosomes for structural and functional differentiation: analysis of the compositions of lipids and proteins in melanosomes and their subfractions. J. Invest. Dermatol., 79, 97–102.

78 Vedralová, E., Duchon, J., and Hach, P.

79

80

81

82

83

84

85

86

87

88

89

(1987) RNA and DNA in melanosomes of hamster melanoma. Pigment Cell Res., 1, 76–80. Borovanský, J. (1997) Detection of metals in tissues, cells and subcellular particles. Sborník Lék., 98, 77–97. Borovanský, J., Duchoň, J., Procházková, B., and Hach, P. (1972) [An attempt to disintegrate melanosomes into protein subunits]. Čas. Lék. Čes., 111, 218–220. Bratosin, S. (1973) Disassembly of melanosomes in detergents. J. Invest. Dermatol., 60, 224–230. Hearing, V.J. and Lutzner, M.A. (1973) Mammalian melanosomal proteins: characterization by polyacrylamide gel electrophoresis. Yale J. Biol. Med., 46, 553–559. Jimbow, K., Jimbow, M., and Chiba, M. (1982) Characterization of structural properties for morphological differentiation of melanosomes: II. Electron microscopic and SDS–PAGE comparison of melanosomal matrix proteins in B16 and Harding-Passey melanomas. J. Invest. Dermatol., 78, 76–81. Seiji, M. and Fitzpatrick, T.B. (1961) The reciprocal relationship between melanization and tyrosinase activity in melanosomes (melanin granules). J. Biochem., 49, 700–706. Menon, I.A. and Haberman, H.F. (1970) Activation of tyrosinase in microsomes and melanosomes from B-16 and Harding-Passey mouse melanoma. Arch. Biochem. Biophys., 137, 231–242. Miyazaki, K. and Seiji, M. (1971) Tyrosinase isolated from mouse melanoma melanosomes. J. Invest. Dermatol., 57, 81–86. Hearing, V.J. (1973) Tyrosinase activity in subcellular fractions of black and albino mice. Nat. New Biol., 245, 81–83. Mufson, R.A. (1975) The tyrosinase activity of melanosomes from the Harding-Passey melanoma: the absence of a peroxidase component in vitro. Arch. Biochem. Biophys., 167, 338–343. Blagoeva, P.M. (1977) Solubilizing effect of Triton X100 on melanosome tyrosinase in hamster pigmented melanoma. Neoplasma, 24, 291–294.

References 90 Kikuchi, A. (1968) Acid phosphatase

91

92

93

94

95

96

97

98

99

100

101

activity in melanosomes of melanocytes. Bull. Tokyo Med. Dent. Univ., 15, 279–294. Novikoff, A.B., Albala, A., and Biempica, L. (1968) Ultrastructural and cytochemical observations on B-16 and Harding-Passey mouse melanomas. The origin of premelanosomes and compound melanosomes. J. Histochem. Cytochem., 16, 299–319. Olson, R.L., Nordquist, J., and Everett, M.A. (1970) The role of epidermal lysosomes in melanin physiology. Br. J. Dermatol., 83, 189–199. Seiji, M. and Kikuchi, A. (1969) Acid phosphatase activity in melanosomes. J. Invest. Dermatol., 52, 212–216. Wolff, K. and Schreiner, E. (1971) Melanosomal acid phosphatase. Arch. Dermatol. Forsch., 241, 255–272. Wolff, K. and Hönigsmann, H. (1972) Are melanosome complexes lysosomes? J. Invest. Dermatol., 59, 170–176. Mufson, R.A. (1974) The subcellular distribution of lysosomal hydrolases in the Harding-Passey melanoma. J. Cell. Physiol., 83, 75–84. Ohtaki, N. (1970) Melanosome and lysosome: I. Lysosomal activity in relation to growth of melanoma. Bull. Tokyo Med. Dent. Univ., 17, 89–102. Kurbanov, C., Abaskina, L.I., Karimova, L.S., and Krivorotova, L.S. (1977) Investigation of tyrosine-α-ketoglutarate transaminase and tryptophane pyrrolase activities in subcellular fractions of skin of guinea pigs. Izv. Akad Nauk Turkmenskoj SSR, Ser. Biol., 72–76. Borovanský, J. and Hach, P. (1999) Disparate behaviour of two melanosomal enzymes α-mannosidase and γ-glutamyltransferase. Cell. Mol. Biol., 45, 1047–1052. Fitzpatrick, T.B. and Quevedo, W.C., Jr (1971) Biological processes underlying melanin pigmentation and pigmentary disorders, in Modern Trends in Dermatology 4 (ed. P. Borrie), Butterworths, London, pp. 122–149. Hill, H.Z. (1992) The function of melanin or six blind people examine an elephant. Bioassays, 14, 49–56.

102 Hu, D.N., Simon, J.D., and Sarna, T.

103

104

105

106

107

108

109

110

111

112

113

(2008) Role of ocular melanin in ophthalmic physiology and pathology. Photochem Photobiol., 84, 639–644. Borovanský, J. (1993) Properties of melanosomes and their exploitation in the diagnosis and treatment of melanoma. Pigment Cell Res., 3, 181–186. Riley, P.A. (1991) Melanogenesis: a realistic target for antimelanoma therapy. Eur. J. Cancer, 27, 1172–1177. Hearing, V.J. (2005) Biogenesis of pigment granules: a sensitive way to regulate melanocyte function. J. Dermatol. Sci., 37, 3–14. Tolleson, W.H. (2005) Human melanocyte biology, toxicology, and pathology. J. Environ. Sci. Health C, 23, 105–161. Ortonne, J.P. and Prota, G. (1993) Hair melanins and hair color. Ultrastructure and biochemical aspects. J. Invest. Dermatol., 101, 82s–89s. Arnaud, J.C. and Boré, P. (1981) Isolation of melanin pigments from human hair. J. Soc. Cosmet. Chem., 32, 137–152. Liu, Y. and Simon, J.D. (2003) Isolation and biophysical studies of natural eumelanins: application of imaging technologies and ultrafast spectroscopy. Pigment Cell Res., 16, 606–618. Novellino, L., Napolitano, A., and Prota, G. (2000) Isolation and characterization of mammalian eumelanins from hair and irides. Biochim. Biophys. Acta, 1475, 295–306. Kushimoto, T., Basrur, V., Valencia, J., Matsunaga, J., Vieira, W.D., Ferrans, V.J., Muller, J., Appella, E., and Hearing, V.J. (2001) A model for melanosome biogenesis based on the puriﬁcation and analysis of early melanosomes. Proc. Natl. Acad. Sci. USA, 98, 10698–10703. Orlow, S.J., Boissy, R.E., Moran, D.J., and Pifko-Hirst, S. (1993) Subcellular distribution of tyrosinase and tyrosinaserelated protein-1: implications for melanosomal biogenesis. J. Invest. Dermatol., 100, 55–64. Van Nieuwpoort, F., Smit, N.P.M., Kolb, R., van der Meulen, H., Koerten, H., and Pavel, S. (2004) Tyrosine-induced

17

18

1 History of Melanosome Research

114

115

116

117

118

119

120

121

122

123

melanogenesis show differences in morphologic and melanogenic preference of melanosomes from light and dark skin types. J. Invest. Dermatol., 122, 1251–1255. Diment, S., Eidelman, M., Rodriguez, G.M., and Orlow, S.J. (1995) Lysosomal hydrolases are present in melanosomes and are elevated in melanizing cells. J. Biol. Chem., 270, 4213–4215. Bhatnagar, V., Anjaiah, S., Puri, N., Darshanam, B.N., and Ramaiah, A. (1993) pH of melanosomes of B16 murine melanoma is acidic: its physiological importance in the regulation of melanin biosynthesis. Arch. Biochem. Biophys., 307, 183–192. Mishima, Y. (1994) Molecular and biological control of melanogenesis through tyrosinase genes and intrinsic and extrinsic regulatory factors. Pigment Cell Res., 7, 376–387. Orlow, S.J. (1995) Melanosomes are specialized members of the lysosomal lineage of organelles. J. Invest. Dermatol., 105, 3–7. Dell’Angelica, E.C., Mullins, C., Caplan, S., and Bonifacino, J.S. (2000) Lysosome-related organelles. FASEB J., 14, 1265–1278. Borovanský, J., Hach, P., Smetana, K., Jr, Elleder, M., and Matouš-Malbohan, I. (1999) Attempts to induce melanosome degradation in vivo. Folia Biol. (Praha), 45, 47–52. Borovanský, J. and Elleder, M. (2003) Melanosome degradation: fact or ﬁction. Pigment Cell Res., 16, 280–286. Kayatz, P., Thumann, G., Luther, T.T., Jordan, J.F., Bartz-Schmidt, K.U., Esser, P.J., and Schraermeyer, U. (2001) Oxidation causes melanin ﬂuorescence. Invest. Ophthalmol. Vis. Sci., 42, 241–246. Sarna, T., Burke, J.M., Korytowski, W., Rózanowska, M., Skumatz, C.M., Zareba, A., and Zareba, M. (2003) Loss of melanin from human RPE with aging: possible role of melanin photooxidation. Exp. Eye Res., 76, 89–98. Jimbow, K., Yamana, K., Akutsu, Y., and Maeda, K. (1988) Nature and biosynthesis of structural matrix protein in melanosomes: melanosomal

124

125

126

127

128

129

130

131

132

133

134

135 136

137

structural protein as differentiation antigen for neoplastc melanocytes, in Advances in Pigment Cell Research (ed. R. Alan), Liss, New York, pp. 169–182. Anonymous (2009) Antibodies speciﬁc to melanocyte-speciﬁc proteins available from the Hearing laboratory. Pigment Cell Melanoma Res., 22, 651. Simon, J.D., Hong, L., and Peles, D.N. (2008) Insights into melanosomes and melanin from some interesting spatial and temporal properties. J. Phys. Chem. B, 112, 13201–13217. Nordlund, J.J., Abdel-Malek, Z., Biossy, R.E., and Rheins, L.A. (1989) Pigment cell biology: an historical review. J. Invest. Dermatol., 4, 53S–60S. Aquaron, R. (1999) Tradition of basic and applied pigment cell research in Marseille. Cell. Mol. Biol., 45, 877–892. Duchoň, J. (1999) Tradition of pigment cell research at Charles University in Prague. Cell Mol. Biol., 45, 893–898. Sangiovanni, G. (1819) Descrizione di un particolare sistema di organi cromoforo espansivo-dermoideo e dei fenomeni che esso produce, scoperto nei molluschi cefaloso. G. Enciclopedico Napoli, 9, 1–13. Westerhof, W. (2006) The discovery of the human melanocyte. Pigment Cell Res., 19, 183–193. Falabella, R. (2009) Vitiligo and the melanocyte reservoir. Indian J. Dermatol., 54, 313–318. Kenney, J.A., Jr (1961) Melanin pigmentation. J. Nat. Med. Assoc., 53, 447–454. Schallreuter, K.U. (2007) Advances in melanocyte basic science research. Dermatol. Clin., 25, 283–291. Gortner, R.A. (1910) Studies on melanin. I. Methods of isolation. The effect of alkali on melanin. J. Biol. Chem., 8, 341–364. Serra, A.J. (1946) Constitution of hair melanins. Nature, 157, 771. Borovanský, J. and Duchoň, J. (1971) Melanoproteins (in Czech). Chem. Listy, 65, 500–528. Nicolaus, R.A. (1968) Melanins, Hermann, Paris.

References 138 Prota, G. (1992) Melanins and

Melanogenesis, Academic Press, San Diego, CA. 139 Swan, G.A. (1974) Structure, chemistry, and biosynthesis of the melanins. Prog. Chem. Org. Nat. Prod., 31, 522–582.

140 Harris, N.B.W., Caddick, M., Kosler, J.,

Goswami, S., Vance, D., and Tindle, A.G. (2004) The pressure–temperature– time path of migmatites from the Sikkim Himalaya. J. Metamorphic Geol., 22, 249–264.

19

wwwwwww

21

2 Classical and Nonclassical Melanocytes in Vertebrates Sophie Colombo, Irina Berlin, Véronique Delmas, and Lionel Larue

2.1 Deﬁnition of Melanogenic Cells

Melanogenic cells produce melanin, a polymer based on tyrosine, in specialized organelles, the melanosomes. This synthesis is catalyzed by an enzyme that converts tyrosine into dopaquinone, tyrosinase (Tyr). In mammals and birds, cutaneous pigment cells are described as “classical melanocytes,” and are involved in skin and hair pigmentation. Melanocytes in other locations, such as the eye, inner ear, meninges, adipose tissue, heart, and, possibly, bone, are described as “nonclassical.” The pigment cell precursor in mammals and birds is called the melanoblast, and the mature pigment cell is called the melanocyte (Figures 2.1 and 2.2). Melanocytes are dendritic cells that produce melanin, the natural pigment of the skin. Melanin is largely responsible for the color of the skin, hair, and eyes, and skin appendages. It plays an important role in protecting skin cells against UV radiation. Melanin is produced in the melanosome, a type of cytoplasmic vesicle related to the lysosome. Some of the quinone intermediate products of melanin synthesis are toxic to cells. The melanosome conﬁnes these compounds, thereby protecting the cell against their harmful effects. Two melanin pigments are synthesized during melanogenesis: eumelanin, which is brown/black in color, and pheomelanin, which is yellow/red in color. The overall color of the appendages and skin results from regulation of the balance between these two types of pigment. Only eumelanin can efﬁciently protect cells against UV. Three enzymes are involved in melanogenesis. The ﬁrst, Tyrosinase (Tyr), plays an essential role in the early stages of melanin synthesis. It hydroxylates l-tyrosine to generate l-3,4-dihydroxyphenylalanine (l-dopa) and then oxidizes l-dopa to generate l-dopaquinone, which spontaneously polymerizes to generate eumelanin. The other two enzymes, tyrosinase-related protein 1 (Tyrp1) and tyrosinase-related protein 2 (Tyrp2, also called dopachrome tautomerase (Dct)), share some similarity with Tyr and ﬁne-tune the synthesis of eumelanin from l-dopaquinone. Pheomelanin is produced from dopaquinone and cysteine. In many mammals, including mice, the agouti signaling pathway regulates this switching of melanin type within cells.

Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

22

2 Classical and Nonclassical Melanocytes in Vertebrates non-classical melanocytes

inner ear

eye

internal organs

RPE

uveal cochlea vestibular meninges harderian others organ gland choroid iris ciliary heart lungs body cartilage, bone gonads, etc classical melanocytes epidermal

dermal

interfollicular follicular niche Figure 2.1 Classical melanocytes are pigmented cells, which (i) are found in the skin (dermis or epidermis), (ii) are involved in skin pigmentation, and (iii) have followed

hair bulb the dorsolateral migratory pathway during development. Melanocytes can be considered as nonclassical if they do not fulﬁll all of these three criteria.

Melanosomes are elliptical or spheroidal organelles containing melanogenic enzymes and cofactors, including proteins of the Tyr family, passing through four stages of maturation [1]. Stage I melanosomes or premelanosomes probably develop from the endoplasmic reticulum [2]. However, the origin of these organelles remains a matter of debate, as Raposo et al. have suggested that stage I melanosomes are actually a type of multivesicular endosome [3]. Stage I melanosomes have an amorphous matrix and internal vesicles formed by membrane invagination. Stage II eumelanosomes have an organized, structured ﬁbrillar matrix, but no active melanin synthesis, whereas melanin synthesis can occur in stage II pheomelanosomes. Stage II eumelanosomes contain Tyr, despite the absence of active melanogenesis. Melanin is deposited on the ﬁbrillar matrix in stage III eumelanosomes and stage IV eumelanosomes are fully melanized, their internal matrix being masked by melanin deposits (reviewed in [4, 5]). Within the skin, melanosomes are transferred to the surrounding keratinocytes, giving the skin its color. Vertebrate pigment cells can be classiﬁed into two types, based on embryonic origin. The cells of one of these types form the retinal pigment epithelium (RPE), which is found in an outer layer of the eye and is derived from an evagination of the developing forebrain, the optic cup. The neural crest-derived pigment cells or melanocytes form the second type of pigment cells. This type of cell includes the pigment cells of the integument and inner ear, the eye and the internal organs. The primary function of these cells is to produce pigment for the coloration of skin, eyes, and appendages (hairs, feathers, scales). Variations in pigment cell

2.1 Deﬁnition of Melanogenic Cells a)

b)

c)

(a) Classical skin melanocytes. X-Gal staining of a transverse section of a Dct:LacZ mouse tail showing melanocytes in the basal layer of the epidermis (Ep) and in the hair follicle (HF). Note the dentricity of the melanocytes. (b and c) Nonclassical

Figure 2.2

heart melanocytes in C57BL/6 mice. Ventral view of whole heart (b) and mitral valve (c). LA, left atrium; LV, left ventricle; MV, mitral valve; RA, right atrium; RV, right ventricle. Scale bars: (b) 1 mm and (c) 100 μm.

development and function are readily identiﬁable and apparent as a particular type of coloration or pattern in vertebrates. Pigment cells are not essential for viability. Mutations affecting their development are, therefore, generally not lethal, but result in obvious changes in the coloring of adult vertebrates. In other vertebrates, the pigment cells are called chromatophores. Chromatophores are pigment-containing, light-reﬂecting cells found in amphibians, ﬁsh,

23

24

2 Classical and Nonclassical Melanocytes in Vertebrates

reptiles, crustaceans, and cephalopods. They are largely responsible for generating skin and eye color in cold-blooded animals, and are generated from the neural crest during embryonic development. Mature chromatophores are grouped into subclasses based on their color under white light: xanthophores (yellow), erythrophores (red), iridophores (reﬂective/iridescent), leucophores (white), melanophores (black/brown), and cyanophores (blue). Melanophores, the equivalent of higher vertebrate melanocytes, contain eumelanin, which is packaged in melanosomes and distributed throughout the cell. In some amphibian species, other pigments are packaged with the eumelanin. For example, a deep-red pigment, pterorhodin, a pteridine dimer that accumulates around the eumelanin core, has been identiﬁed in the melanophores of some frogs. Melanophores display specialized bidirectional and coordinated translocation of the melanosomes within the cytoplasm. Melanophores develop from the neural crest, and are most abundant in the dermal and epidermal layers of the skin, in which the intracellular distribution of the pigment has a signiﬁcant effect on the color of the animal. Pigment transport is dependent on an intact cytoskeleton and motor proteins associated with cytoskeletal components. Some species can change color rapidly, through mechanisms involving pigment translocation and the reorientation of reﬂective plates within chromatophores. Such mechanisms are often used as a type of camouﬂage. Certain vertebrates, including chameleons, generate this effect by producing cell signaling molecules, such as hormones or neurotransmitters, in response to changes in mood, temperature, stress, or visible changes in local environment. Zebraﬁsh and amphibians have three types of pigment cell (melanophores, xanthophores, and iridophores), and medaka (Japanese killiﬁsh) have a fourth type, leucophores. The distribution of the different types of chromatophores is the major determinant of the pattern of coloration in adult ﬁsh and amphibians.

2.2 Distribution and Function of Melanogenic Cells

Melanocytes are a relatively recent development on the evolutionary timescale. They appeared after the formation of the fourth germ layer or neural crest. They have since evolved to execute various functions, depending on their location, but they retain common signature functions, to which melanin synthesis makes an essential contribution. The best-known function of melanocytes is the one of classical melanocytes, which, due to their location in the skin, are the most apparent and widely studied. These cells control the pigmentation of the individual, tanning, and protection against UV rays. The melanogenic cells in the eye are also highly apparent. Histological analysis rapidly revealed that the cells of the RPE differ in shape from skin or uveal melanocytes. Melanocytes and/or melanin pigment were subsequently identiﬁed within the body, in the inner ear, heart, brain, and adipose tissue. The functions and the origins of these nonclassical melanocytes remain unclear.

2.2 Distribution and Function of Melanogenic Cells

2.2.1 Classical Melanocytes

Classical melanocytes are found in the epidermis and dermis. In the epidermis, they are located either in its basal layer or in the hair follicles. The cells surrounding the epidermal melanocytes are keratinocytes, a type of epithelial cell, whereas dermal melanocytes are surrounded mostly by ﬁbroblasts. Epidermal melanocytes can transfer melanin to the surrounding cells, whereas dermal melanocytes cannot. The distribution of melanocytes may differ with species. In humans, melanocytes are mostly located in the basal layer of the epidermis, whereas, in rodents, they are found mostly in hair follicles. Rodent melanocytes are found in the epidermis of nonhairy skin (tail, ears, nose and footpad) and in the dermis of the pinna of the ears. The skin is the main barrier to the external environment. Melanin synthesis in the skin evolved as a protection against the harmful ionizing radiation of sunlight. The enzymes involved in melanin synthesis are known to modulate the levels of calcium and reactive species [6–8]. Skin melanocytes protect the body from the harmful effects of light and heat, by producing melanin. Skin phototype, corresponding to skin color and ease of tanning, reﬂects the degree of pigment production in humans and is the best indicator of skin cancer risk in the general population. Individuals with lightly pigmented skin have a much higher risk (15–70 times higher) of skin cancers, including melanomas, than individuals with darker skin [9, 10]. The melanocortin-1 receptor (MC1R) is a major regulator of skin pigmentation phenotype and the gene encoding this protein has thus been identiﬁed as a potential susceptibility gene for melanoma [11, 12]. UV radiations are harmful to human skin because they generate several types of cellular damage, including oxidative damage and two major types of DNA damage: cyclobutylpyrimidine dimers and 6,4-photoproducts, which play a major role in UV carcinogenesis [13]. Its seems that DNA damage/repair itself induces skin pigmentation [12, 14, 15]. 2.2.1.1 Melanocytes in the Epidermis Epidermal melanocytes constitute the second largest cell population (about 5%) in the human epidermis after keratinocytes, which account for about 90% of epidermal cells. They synthesize the eumelanins and pheomelanins that give the skin its color and protect it against UV light. They fulﬁll various roles, including protecting the skin from DNA damage or oxidative stress and immune responses to various stimuli. 2.2.1.1.1 Melanocytes in the Hair Follicle The melanocytes in hair follicles are involved in both hair pigmentation and the elimination of toxic byproducts of melanin synthesis [16]. The bulge region of the hair follicle serves as a reservoir of melanocyte stem cells for the skin and hairs. These stem cells give rise to new melanocytes populating the bulb region of the hair follicle at each hair cycle.

25

26

2 Classical and Nonclassical Melanocytes in Vertebrates

2.2.1.1.2 Interfollicular Melanocytes Melanin has many roles. It protects against the DNA damage and free radicals induced by UV radiation. It also slows vitamin D production in response to UV-B irradiation and protects against the UV-A-mediated breakdown of folate, which is essential for nucleotide synthesis. It has also been suggested that pigmentation may be involved in adaptation to ambient temperature and sexual selection for certain species. Epidermal melanocytes are surrounded by keratinocytes, which they contact via their dendrites, in a ratio of about 30 keratinocytes per melanocyte. The epidermal melanocyte and its surrounding keratinocytes form the so-called “epidermal melanin unit.” Melanosomes are transferred from the melanocytes to the keratinocytes, in which their distribution is modiﬁed by UV radiation such that they form a cap-like structure covering the side of the nucleus exposed to sunlight [17, 18]. They protect epidermal cells from damage by limiting the penetration of UV rays through the epidermal layers and by scavenging reactive oxygen species (ROS) generated in response to UV exposure [19, 20]. Melanocytes express MC1R, which regulates the quality and quantity of melanin production. MC1R is controlled by the agonists melanocyte-stimulating hormone (MSH) and adrenocorticotropic hormone (ACTH) [21], which stimulate the eumelanin synthesis pathway. This receptor is also controlled by the antagonist agouti signaling protein (ASIP), which activates the production of pheomelanin [18, 22]. By responding to paracrine factors, such as α-MSH, epidermal melanocytes can repair DNA more efﬁciently by nucleotide excision repair [23, 24], and also combat oxidative stress, because α-MSH decreases hydrogen peroxide generation in response to UV irradiation [23]. These consequences are thought to decrease the genotoxic effects of UV irradiation, enabling melanocytes to survive with a stable genome. The maintenance of melanocyte survival in the skin is essential to optimize photoprotection and to prevent photocarcinogenesis [18]. The skin is continually exposed to microorganisms. The epidermis has no vascular circulation, minimizing the likelihood of toxic chemicals, bacteria, or fungi penetrating the stratum corneum and entering the bloodstream. The epidermis protects itself by producing defensins, a group of proteins with antimicrobial properties. It also contains Toll-like receptors, which recognize invading organisms and elicit a host response. The reactive quinone intermediates generated during melanin biosynthesis also have potent antimicrobial/antifungal properties. Consistent with this ﬁnding, fungal dermatitis is more prevalent among individuals with fair skin than among those with dark skin [25]. Melanocytes are also phagocytic cells involved in the inﬂammatory response. Indeed, α-MSH is known to suppress inﬂammation (particularly by inhibiting the activation of nuclear factor NF-κB). The pigment system responds to almost all inﬂammatory events in the epidermis, generally by increasing melanin synthesis (postinﬂammatory hyperpigmentation), or less often, by decreasing melanin synthesis (postinﬂammatory hypopigmentation). An example of postinﬂammatory hyperpigmentation is provided by tanning after sun exposure. UV rays damage the epidermis and sunburn is the inﬂammatory response to this injury. This process is mediated in part by α-MSH, which induces melanin production, result-

2.2 Distribution and Function of Melanogenic Cells

ing in darker skin that is more resistant to subsequent sunburn. Further functions of α-MSH include the stimulation of DNA repair and downregulation of the immune response of the epidermis, possibly preventing the occurrence of autoimmune disorders, such as lupus erythematosus [18]. Melanocytes produce and react to many cytokines and growth factors, thereby acting as sentinels and active players in the immune system of the skin [2]. Melanocytes can present and process antigens, eliciting a T cell proliferative response [26], with interferon-γ inducing the expression of major histocompatibility complex class II (MHC II) antigens [27]. Melanocytes also interact with Langerhans cells, the principal type of antigen-presenting cell in the skin [18, 28]. Melanocytes probably also have an endocrine/sensory function, regulating the skin immune response. The cutaneous pigment system can serve as an important stress response element, through the effects of corticotrophin-releasing hormone (CRH), catecholamines, acetylcholine (ACh), pro-opiomelanocortin (POMC)derived peptides, and steroid hormones [29–31]. CRH has been shown to inhibit NF-κB (a proinﬂammatory molecule) in melanocytes by producing POMC and may therefore serve as a feedback mechanism for the self-restriction of inﬂammatory responses in the skin [32]. Lipocalin-type prostaglandin D synthase (L-PGDS) has been identiﬁed as a melanocyte marker, due to its speciﬁc expression in mouse and human epidermal melanocytes, but not in other skin cells [33]. L-PGDS isomerizes prostaglandin H2 (PGH2) to generate prostaglandin D2 (PGD2) and functions as an intercellular chaperone for lipophilic ligands, such as thyroid hormones, retinoic acid, bilirubin and biliverdin [34]. PGD2 has pleiotropic effects through its speciﬁc G-proteincoupled receptors, DP1 and DP2, both of which are expressed in human epidermal melanocytes. Mitf-M regulates L-PGDS gene transcription in melanocytes. L-PGDS and PGD2 regulate sleep and nociception. Melanocytes also produce β-endorphin, an endogenous opioid, from POMC, which, together with MSH and ACTH and opioid receptors, is located in nuclei active in sleep regulation. It has therefore been suggested that melanocytes may be involved in regulating sleep/waking behavior [35]. There is also evidence to suggest that a melanocyte-derived factor, such as L-PGDS, may be involved in regulating the central chemosensor controlling respiratory rhythm, as opioids are respiratory depressants and epidermal melanocytes produce opioids [35]. 2.2.1.2 Melanocytes in the Dermis Dermal melanocytes are considered to be classical melanocytes because they are involved in skin pigmentation. However, they do not transfer melanin to surrounding cells. They do not interact with keratinocytes, instead interacting with ﬁbroblasts and other cell types, which may instruct them. Dermal melanocytes are present in very small numbers in human adult skin and little is known about their function. Their interaction with the surrounding ﬁbroblasts seems to regulate their melanin production. Fibroblasts have been shown to stimulate melanocytes to produce Tyr [36], thereby increasing melanocyte survival and melanin synthesis after the UV irradiation of reconstructed skin [37]. By contrast, Hedley et al. [38]

27

28

2 Classical and Nonclassical Melanocytes in Vertebrates

showed that ﬁbroblasts decreased pigmentation levels in reconstructed skin models. Yamaguchi et al. [39] showed that Dickkopf1 (DKK1), an inhibitor of Wnt/β-catenin signaling, was produced in larger amounts in palmoplantar dermal ﬁbroblasts than in nonpalmoplantar dermal ﬁbroblasts. DKK1 inhibits melanocyte function and growth, by regulating β-catenin and microphthalmia-associated transcription factor (Mitf) [40]. In addition to modulating melanocyte melanin production, DKK1 also affects keratinocytes, increasing skin thickness and decreasing the transfer of melanosomes from melanocytes to keratinocytes. 2.2.1.3 Melanophores in Lower Vertebrates Frog melanophores of various morphological types are present in both the epidermis, which is populated mostly by keratinocytes, and the dermis, in which ﬁbroblasts predominate. Amphibian and ﬁsh melanophores perceive chemical stimuli and play an important role in rapid color changes, through the dispersion or aggregation of melanosomes, and neurogenic changes in pigmentation [41]. These modiﬁcations of pigmentation are essential for the survival of these animals in predator-rich environments. Unlike the melanocytes of mammals, amphibian melanophores can cross the basal lamina to reach the dermis, where they transfer melanin to dermal ﬁbroblasts [42]. 2.2.2 Nonclassical Melanocytes

Nonclassical melanocytes can be found in various locations of the body, including the visceral organs. They do not usually contain large amounts of melanin in healthy individuals. However, there are key exceptions to this rule. The abdomen of birds, including quail, chicken, ostrich, and zebra ﬁnch, contains pigmented cells, and the spleen and liver of amphibians contain pigmented macrophages. Nonclassical melanocytes are more commonly found in the eye, inner ear, heart, brain, and adipose tissue. 2.2.2.1 Melanocytes of the Eye The eye contains two types of pigment cells: the uveal melanocytes and the RPE cells. RPE cells play a crucial role in retinal development and function, thus controlling visual acuity, whereas uveal melanocytes provide the color of the iris and are responsible for uveal melanoma development. Other melanocytes are found in the Harderian gland of vertebrates with nictitating membranes, which may play a role in photoprotection. Photoreception, in general, and in the eyes, in particular, is commonly associated with dark pigments, such as melanins, because of their ability to absorb light. Such pigments are found in all functional eyes, even the most primitive. By screening out light from one direction only, they protect the photoreceptor cells from excessive exposure to light. The amount of light reaching the photoreceptors is also regulated by the pupil, the diameter of which varies with light intensity. The photoreceptive retina in vertebrates is covered by a layer of choroidal melanocytes and a specialized pigmented monolayer, the RPE. The RPE

2.2 Distribution and Function of Melanogenic Cells

cells and their melanosomes do not seem to be renewed, whereas the uveal melanocytes and their melanosomes are produced continually, throughout the life of the individual. 2.2.2.1.1 RPE The RPE plays a key role in visual acuity, for which melanin production does not appear to be required. In vertebrates, the cells of the RPE are derived from the multipotent optic neuroepithelium. They develop in close proximity to the retina, and are indispensible for eye organogenesis and vision. RPE cells are cuboidal cells that contact the outer segments of the photoreceptors on their apical side and rest against the basal membrane, known as Bruch’s membrane, separating the RPE from the choroid, at their base. During embryogenesis, RPE cells are involved in the formation of the ciliary body and iris, and the control of optic ﬁssure closure. They also inﬂuence retinal neurogenesis and ganglion cell projections, and have been implicated in regulation of the choroidal vasculature (for review, see [43, 44]). In adults, RPE cells provide photoreceptors with nutritional support, build up a blood–retina barrier and process retinol and retinoids, controlling ion ﬂow and oxidative damage, and removing the fragments of photoreceptor outer segments shed from the distal end of the photoreceptor by phagocytosis [45]. The melanin in the RPE protects the neural retina against ROS, thereby preventing age-related macular degeneration [18]. RPE melanin is essential for the development of the neural retina. When it is absent or present at low levels, the retina develops abnormally (e.g., in albino mammals) and many temporal retinal axons that should remain uncrossed at the optic chiasm cross inappropriately to innervate the contralateral hemisphere. The central retina is also underdeveloped and there is a rod cell deﬁcit of about 30% (for a review, see [46]). These retinal abnormalities result in signiﬁcant visual impairment. Albino birds do not display the same retinal abnormalities, probably because, unlike mammals, they have a cone-dominated retina with few rods. The synthetic pathway of melanin is the same in the skin and in the RPE, but unlike epidermal melanocytes, which produce melanosomes continuously, RPE cells are thought to produce melanosomes only during the prenatal period. Both prenatal RPE cells and skin melanocytes present stage I–IV melanosomes. Melanogenesis can occur only if premelanosomes are present, and Tyr and other melanogenic proteins are synthesized. The adult RPE contains Tyr, but not the structural protein Pmel17, which is essential for the formation of premelanosomes in melanocytes. Biesemeier et al. [47] showed that melanogenesis can occur within multivesicular and multilamellar organelles in amelanotic RPE cells in vitro, without the formation of typical premelanosomes. Tyrp1 and Pmel17 are not detectable in these cells and classical stage I–IV melanosomes are also absent. 2.2.2.1.2 Uveal Melanocytes The uvea is the pigmented, highly vascularized middle layer of the eye. It is divided, from anterior to posterior, into the iris, ciliary body, and choroid. The pigment is produced and held in numerous dendritic melanocytes, similar to

29

30

2 Classical and Nonclassical Melanocytes in Vertebrates

normal dermal melanocytes. It has been suggested that uveal melanin reduces the likelihood of uveal melanoma by protecting melanocytes from oxidative stress. The role of the melanin in uveal melanocytes in protection against UV remains a matter of debate [48, 49]. Choroid The choroid, a layer of highly vascularized tissue surrounding the eye, is essential for normal retinal function. The melanocytes in this structure are localized in the choroidal stroma and suprachoroid. They are pigmented and may be star-shaped, fusiform, round, or oval, with a central nucleus. Their function remains unclear, but the ﬁbrillar structures observed in their cytoplasm may be involved in maintaining capillary tonus. There is no functional contact between the dendrites of the choroidal melanocytes and the other cell types present in uveal tissue. These melanocytes remain quiescent and no longer have the capacity to produce melanin [49]. Choroidal melanin is thought to protect the choroid from oxidative damage due to the stress arising from its location in the highly vascularized posterior segment of the eye. Iris The iris is a small structure consisting of connective tissue and muscle, with a central opening called the pupil. Eye color, reﬂecting the pigmentation of the iris, depends on the total amount of melanin in the epithelial cells and stromal melanocytes and the relative proportions of the two types of melanin. Brown irises contain abundant melanocytes and melanin in the anterior border layer and stroma, whereas these layers contain very little melanin in blue irises. This pigmentation is essential in the control of pupil aperture (thus controlling the amount of light entering the eye). The pigmentation of the iris appears to be correlated with the incidence of uveal melanoma and age-related macular degeneration, for which it is considered a risk factor [48, 49]. Ciliary Body The ciliary body is the circumferential tissue within the eye consisting of the ciliary muscle and ciliary processes and coated with a double layer, the ciliary epithelium. The outer layer is highly pigmented and continuous with the RPE. These cells constitute the iris dilator muscle, which plays an important role in light accommodation, by controlling pupil aperture. 2.2.2.1.3 Harderian Gland The Harderian gland is found in the orbit of the eye in vertebrates (reptiles, amphibians, birds, and mammals) with a nictitating membrane. Dendritic cells containing melanin granules have been found in the interstitial tissue of the Harderian gland, which is pigmented. Two types of melanocytes, with and without the various developmental stages of melanin granules, have been found in this gland in mice. Cells with developing granules were found to have more dendrites and to contain a larger number of cytoplasmic organelles. The other cells were ellipsoidal or elongated and contained few cytoplasmic organelles and a large number of fully pigmented melanin granules, but no developing granules. The Harderian gland macrophages contain fully pigmented melanin granules of

2.2 Distribution and Function of Melanogenic Cells

various sizes, suggesting that they engulf some of the melanocytes. The numbers of mature melanocytes in the Harderian gland probably varies with retina pigmentation. For example, gerbil strains with pink eyes also have Harderian glands lacking melanocytes. The role of melanocytes in the Harderian gland is unclear, but may be related to photoprotection. The nictitating membrane is also pigmented in many species, with melanocytes present around the duct and its opening [50]. 2.2.2.2 Melanocytes of the Inner Ear Corti was the ﬁrst to demonstrate the presence of melanocytes in the inner ear, in 1851. The distribution of melanocytes in the inner ear differs between species. These cells are mostly concentrated in the cochlea and the vestibular organ. Melanocytes are abundant in the human inner ear, but are rare in the ears of lower vertebrates. Patients with lacking inner ear melanocytes or with poor or no inner ear melanocyte function, such as those suffering from Waardenburg syndrome or Harada’s disease, display hearing loss and problems with balance. This suggests that melanocytes have important functions in the cochlea and the vestibular labyrinth. Inner ear melanocytes have also been shown to play an important role in animal models, such as viable dominant spotting mice or white spotting rats, which have a mutation in the c-Kit gene (KitW), and in black-eyed white Mitfmi-bw homozygous mice and Mitfmi-vga9 homozygous mice. All these animals have coat color phenotypes due to a lack of melanocytes and are deaf. In addition, mice homozygous for the weak allele, Mitfvit, gradually lose their melanocytes after birth, and thus display hair graying and age-dependent hearing loss, indicating a role for Mitf and melanocytes not only during inner ear development, but also in inner ear function after development has been completed. 2.2.2.2.1 Cochlea The cochlea is the organ responsible for hearing in mammals. Cells containing melanin are present within the stria vascularis and the modiolus. These cells play important roles in the development and function of the cochlea, by regulating endocochlear potential and preventing noise-induced damage (for review, see [51]). The endolymph – the extracellular ﬂuid in the membranous labyrinth of the inner ear – is excreted by the stria vascularis, which consists of three layers of cells: the marginal, intermediate, and basal cell layers. The intermediate layer cells in the stria vascularis of the mammalian cochlea are melanocytes that contain and synthesize melanin. Two types of cochlear melanocytes have been described: light intermediate cells that contain premelanosomes, are positive for the dopa reaction, and capable of melanin synthesis, and dark intermediate cells corresponding to degraded melanocytes incapable of melanin synthesis. In the modiolus, the melanocytes are located in the perivascular and perineural spaces. These cells are closely associated with the spiral modiolus artery. Cochlear melanocytes are required for the normal development and function of the cochlea, as mammals with pigmentation defects display profound cochlear defects. Viable dominant spotting mice (KitWv) and white spotting rats (KitWs) have no cochlear melanocytes and

31

32

2 Classical and Nonclassical Melanocytes in Vertebrates

display alterations in the development and function of the stria vascularis, including changes in endocochlear potential. However, melanin itself is not essential for normal hearing function. The absence of melanin has no effect on age-related cochlear degeneration [52]. Furthermore, albino mammals, which have melanocytes but no pigment due to a lack of Tyr or related enzyme function, have normal hearing. However, albino cochleas are more susceptible to the ototoxic effects of high-intensity noise than pigmented cochleas [51]. This may be accounted for by the ability of melanin to scavenge the ROS, generated after exposure to noise. Melanin formation in the inner ear seems to be catalyzed by a tyrosine hydroxylase rather than by Tyr [53]. Tyr and Tyrp1 have not been found in cochlear extracts from the gerbil, but these extracts do contain tyrosine hydroxylase with a higher afﬁnity for l-tyrosine, and this enzyme is independent of l-dopa. Strial intermediate cells continually undergo mitosis. Exposure to noncontinuous impulse-type noise or broadband white noise leads to strong hyperpigmentation of the stria vacularis, with some melanosomes being transferred to marginal cells. An increase in prostaglandin levels after noise exposure may lead to an increase in the mitosis rates of intermediate cells [51]. Intermediate cells generate a positive endocochlear potential and secrete K+. The + K channel at the plasma membrane may be involved in the mitosis of intermediate cells. Mice lacking these K+ channels are deaf. Melanocyte-deﬁcient mice display measurable changes in the ionic composition of the endolymph, which bathes the mechanosensory hair cells [54]. Normal endolymph contains high levels of K+ ions, and low levels of Na+ and Ca2+ ions. Cochlear melanocyte deﬁciencies are associated with a low K+ concentration in the endolymph, affecting the capacity of hair cells to respond to the sound-induced motion of the endolymph [55]. The melanin in cochlear melanocytes may also serve as a biological reservoir for Ca2+ and may be involved in Ca2+ regulation in the inner ear, as it has a high afﬁnity for divalent cations, such as Ca2+ [56, 57]. Variations in endolymph Ca2+ concentration are known to affect the function of hair cells. Albino animals, which have amelanotic melanocytes, also have low concentrations of Ca2+ in the endolymph. However, the molecular mechanisms by which these melanocytes exert their functions are unknown. The glutathione S-transferase (GST) α4 (Gsta4) gene has recently been shown to be expressed speciﬁcally in melanocytes of the stria vascularis, but not in other tissues harboring melanocytes [58]. This suggests that cochlear melanocytes play a role independent of their melanogenic function. GSTs carry out important functions in the detoxiﬁcation processes of many tissues. Other GSTs, including Gsta1/2, Gstm1/2, and Gstp1, are expressed in the lateral wall of the cochlea, which includes the melanocyte-containing stria vascularis. Gstp1 seems to be expressed in the intermediate cells and may therefore colocalize with Gst4. Alpha GSTs play a role in defense against the intracellular lipid peroxidation caused by the reaction of various lipids with ROS. Gsta4-4 (Gsta4 homodimer) is the major enzyme catalyzing the conjugation of glutathione with 4-hydroxynonenal, one of the end products of lipid peroxidation. Gsta4 in melanocytes of the stria vascularis may act as an antioxidant in defenses against the hearing loss induced by loud noises and ototoxic drugs, which have been reported

2.2 Distribution and Function of Melanogenic Cells

to induce oxidative stress through the production of ROS. This stress results in deafness, due to the injury of hair cells in the cochlea. ROS are also inevitably produced during normal aerobic metabolism. Blood vessels in the stria vascularis supply oxygen and may be a source of oxidative stress. The distribution of GSTs may facilitate the detoxiﬁcation of harmful substances at the boundaries between blood vessels and the tissues of the inner ear. Cochlear melanogenesis is induced by exposure to noise. Pigmented cochleas are less susceptible to noise partly because melanin, as an energy transducer, works as a ROS scavenger. The speciﬁc expression of Gsta4 in cochlear pigment cells may therefore be of particular importance, with this molecule acting as an antioxidant and its levels being correlated with cochlear melanogenesis. 2.2.2.2.2 Vestibular Organ Melanocytes are also present in the vestibular labyrinth of the inner ear [59]. In viable dominant spotting mice, which have profound defects of the stria vascularis, the vestibular labyrinth appears to be pigmented in approximately 20% of animals. The melanocytes of the vestibular labyrinth have been shown to display the characteristics of epidermal melanocytes in culture, with the presence of melanosomes and the expression of genes associated with pigmentation, such as the Melan-A, Tyrp1, and Tyr genes. Their function remains unclear. It has been speculated that they could play a role in balance perception, as melanocyte defects have been found to be accompanied by vertigo. The vestibular melanocytes are located in the dark cell areas of the vestibular labyrinth. They form a continuous pigmented layer covered by a secretion epithelium consisting of epithelial dark cells (EDCs) and are presumably in contact with neighboring subepithelial capillaries. These cells have large numbers of cytoplasmic processes and melanosomes at various stages. Gap junctions are found between melanocytes and between melanocytes and EDCs. These junctions may be involved in communication and the endolymph–perilymph transport of molecules. They may also play a role in the regulation of local Ca2+ homeostasis. Changes in the morphology and activity of vestibular inner ear melanocytes (VIEMs) have been observed in pathological changes to the vestibular labyrinth, during experimental hydrops, or in the presence of gentamicin, for example. These changes suggest that VIEM may react to stress conditions in the inner ear. EDCs are involved in endolymphatic K+ circulation. By analogy, melanocytes may therefore be involved in K+ transport or in the constitution of the endolymph in the vestibular labyrinth, in the same way that the melanocytes of the stria vascularis have K+ channels in their membrane and contribute to plasma membrane potential. However, no such K+ channels were found in the dark cell areas and vestibular function remains normal in mice lacking these channels. 2.2.2.3 Melanocytes of the Heart Melanocytes are found in many locations in the heart, including the valves and septa. These melanocytes are dependent on and produce proteins of the classical melanocyte signaling pathway. The level of pigmentation in the heart reﬂects that

33

34

2 Classical and Nonclassical Melanocytes in Vertebrates

in the skin. The function of cardiac melanocytes remains unclear, because these cells do not seem to be essential for normal heart function. They may play a role in atrioventricular (AV) valve function, by modulating the viscoelastic properties of these valves. They also contribute to the triggering of arrhythmia, by regulating calcium and ROS levels. It has been known for some time that the neural crest contributes to the formation of the normal heart, particularly in terms of the septation of the outﬂow tract and the formation of the venous pole. However, melanocytes have only recently been found in the heart [60]. It has been shown in vitro that cardiac neural crest cells (NCCs) form melanocytes. Furthermore, quail-to-chicken transplantation experiments have demonstrated that cranial NCCs form pigmented cells on all the major arteries, including those at the base of the heart. Pigmented cells were also found in the hearts of mutant mice from a line derived from a large-scale N-ethylN-nitrosourea mutation screen. Several groups have investigated the distribution of pigmented cells in the heart [60–63]. l-Dopa-positive melanocytes containing Dct and Tyrp1 are found within the AV mitral (left side) and tricuspid (right side) valve leaﬂets, and are also present in the chordae tendineae, which extend from the distal edge of the leaﬂet to the papillary muscle. They are found in the aortic semilunar valve leaﬂets and in the AV septa. They are present at the apex of the interventricular septum, just below the endocardial layer. They are also present on the dorsal surface of the medial wall of the right atrium, close to the conﬂuence of the coronary sinus and the superior vena cava, and on the outer medial wall of the right atrium. These cells are detected along the right interatrial septum near the foramen ovale and toward the septal leaﬂet and the aorta. Levin et al. [62] also found melanocytes in the pulmonary veins of mice and humans, although those present in human pulmonary veins were not pigmented, probably because they did not express the Tyrp1 gene. Very small numbers of melanocytes have recently been found in the ductus arteriosus and ligamentum arteriosum of mice [64]. They are never observed in the pulmonary valve leaﬂets, muscular wall of the interventricular septum or papillary muscle. Melanoblasts are ﬁrst detected in the heart at embryonic day (E) 12.5, in AV endocardial cushions [61], and prevalvular cushions of the AV region of E13 embryonic hearts are capable of producing pigmented cells in culture [60]. The cardiac melanocytes express markers of the classical melanocyte lineage, such as Dct, Sox10, Mitf, Tyr, Tyrp1, and Pax3. They also differ from cardiomyocytes in terms of their morphology. They have large numbers of caveolae along the plasma membrane and some cells contain mature melanosomes [62]. No melanin appears to be transferred from cardiac melanocytes to the surrounding cells [63]. A cluster analysis of global gene expression suggested that Dct-expressing cells were more closely related to atrial myocytes than to dermal melanocytes. These two cell types have similar calcium-handling proteins (such as phospholamban and ryanodine receptor, type 2) and have some voltage-gated ion channels in common (e.g., the pore-forming subunit of L-type calcium channels and cardiac voltage-dependent sodium channels). However, Dct-positive cells in the heart display no signiﬁcant expression of some of the signature genes of cardiomyocytes,

2.2 Distribution and Function of Melanogenic Cells

such as those encoding cardiac actin and troponins. Thus, Dct-expressing cells in the heart can be distinguished from atrial myocytes and dermal melanocytes on a molecular basis [62]. The distribution of melanocytes in the heart is similar in mice lacking Dct and wild-type mice. Dct is therefore not required for the migration and survival of melanocytes in the heart. The level of pigmentation in the heart seems to be correlated with coat color in mice. Cardiac melanocytes were not found in the spotting mutants Ednrbsl/sl and KitWv/Wv, which have very few melanoblasts and are almost completely white [61], or in the Mitfvga9/vga9 mutants, which are white due to a lack of melanocytes; bcat* mice also have very few cardiac melanocytes and present hypopigmentation [63]. Conversely, large numbers of cardiac melanocytes are present in Tyr::N-rasQ61K transgenic mice, which produce the oncogenic N-Ras in melanocytes, or in mice that overproduce endothelin 3 (Edn3) or hepatocyte growth factor [61, 63]. In mice overproducing Edn3, ectopic melanocytes have even been found in the pulmonary valve and outﬂow tract. These results indicate that cardiac melanocytes are dependent on the signaling molecules known to be required for correct skin melanocyte development, suggesting that they may originate from the same precursor population. Cardiac melanocytes have not been found in zebraﬁsh or frog, but are present in quail, suggesting an association between the presence of cardiac melanocytes and four-chambered hearts. The function of cardiac melanocytes remains unclear. These cells are not essential for gross heart morphogenesis and physiology, as mice lacking melanocytes (e.g., KitWv or Mitfvga9 mice) and mice with abundant pigmentation (e.g., Tyr::NrasQ61K mice) in the valves and septa of the heart live well into adulthood, with apparently normal hearts. These cells may therefore play a more subtle role that becomes critical in stress conditions. They may be involved in AV valve development from the endocardial cushions, as they arrive in the heart in these cushions. Melanoblasts have been shown to produce and secrete metalloproteases and may thus participate in valve remodeling. This function may continue into adulthood, during which both melanocytes and interstitial cells may be involved in the maintenance of tissue homeostasis and may affect the mechanical properties of the AV valves. Cardiac melanocytes increase the stiffness and storage of mouse AV valves, thereby affecting their viscoelastic properties, and are important for the correct functioning of these valves in the body [65]. Quasistatic and nanodynamic mechanical analyses of the leaﬂets of the mouse tricuspid valve have indicated that the mechanical properties of the leaﬂet vary with the degree of pigmentation. Melanocytes may be stiffer than the endothelial cells overlying the leaﬂet or the surrounding extracellular matrix. However, it remains unclear whether it is the melanocytes themselves or the presence of melanin that contributes to the mechanical properties of the leaﬂets. It is also possible that variations in stiffness result from the inﬂuence of melanocytes on the surrounding extracellular matrix. As melanocyte precursors reach the heart early in valve development and persist into adulthood, dysfunctions affecting these precursors might lead to valve abnormalities. Indeed, cardiac melanocytes are involved in the triggering of atrial arrhythmia [62]. Atrial ﬁbrillation is the most common form of cardiac arrhythmia. It is often initiated by ectopic beats arising from the pulmonary veins and atrium. The

35

36

2 Classical and Nonclassical Melanocytes in Vertebrates

dysregulation of intracellular calcium and reactive species levels has been described in patients with atrial ﬁbrillation. Dct is involved in the regulation of these concentrations in melanocytes. Mice with no Dct in their cardiac melanocytes have been shown to be more susceptible than wild-type mice to atrial arrhythmogenesis, in the absence of other cardiac electrophysiological alterations and structural abnormalities. By contrast, mice lacking cardiac melanocytes (KitW/Wv mutants) display no atrial arrhythmia. Cardiac melanocytes express adrenergic and muscarinic receptors and interact with autonomic nerves (sympathetic and cholinergic nerves). They can therefore respond to autonomic efﬂux, which may contribute to arrhythmia. The treatment of mice lacking melanocytes with muscarinic agonists or of mice with melanocytes with β-adrenergic antagonists reduces atrial arrhythmia. The treatment of mice lacking Dct with antioxidants signiﬁcantly decreases the frequency of atrial arrhythmia, conﬁrming the antioxidant role of Dct and suggesting a role for ROS in triggering arrhythmia in the absence of Dct. Cardiac melanocytes are excitable and form electrical connections with neighboring cardiomyocytes. In the absence of Dct, cardiac melanocytes display prolonged repolarization, with early after depolarization and frequent calcium oscillations, conﬁrming the role of Dct in calcium handling and that of intracellular calcium dysregulation in arrhythmogenesis. Cells capable of synthesizing melanin may be present in the heart and pulmonary veins to buffer calcium and reactive species. However, during pathologic processes, the capacity of these cells to bind free radicals and calcium may be changed or their binding sites saturated, transforming them into initiators of triggered activity and cardiac arrhythmia. Melanin-synthesizing cells may also be involved in maintaining the normal balance of oxidative species in the myocardium. 2.2.2.4 Melanocytes of the Brain and Neuromelanins Melanin-producing melanocytes are found in the sympathetic cephalic ganglia and leptomeninges, and along cerebral capillaries. In the leptomeninges, these cells principally cover the ventrolateral surfaces of the medulla oblongata. These dendritic cells contain melanosomes, observed as single, membrane-bound granules and displaying all stages of melanization, thus conﬁrming that they are indeed melanocytes and not macrophages. At other locations in the brain, the meninges contain only isolated pigmented cells [66]. Brain melanocytes may have a neuroendocrine and detoxiﬁcation function. Epidermal melanocytes produce L-PGDS, a potent inducer of sleep, and produce opioids that regulate respiratory rhythm. It is thus possible that brain melanocytes produce such molecules in vivo and regulate these mechanisms. Neuromelanin, a brain-speciﬁc pigment, is found primarily in catecholaminergic neurons (which produce norepinephrine and dopamine, but not epinephrine) in the substantia nigra and locus coeruleus [67], and accumulates with age [68]. Neuromelanin has a spherical architecture and is composed of a pheomelanin core covered with eumelanin; it also contains aliphatic compounds and peptides [69]. It is formed from the oxidation products of dopamine and cysteinyldopamine in the substantia nigra, and is derived from norepinephrine metabolism in the

2.3 Embryonic Development of Melanogenic Cells

locus coeruleus. It is abundant in the human brain and present in smaller amounts in some nonhuman primates, but it is not found in the brains of many lower species. Neuromelanins play a critical role in protecting neurons against toxic ROS and metals, such as iron [70–72]. Neuromelanins produced from dopa and cysteinyldopa have recently been found in many other types of neuron, almost everywhere in the brain. They also accumulate with age and may play an essential role in reducing toxicity in these tissues [73]. In Parkinson’s disease, more dopaminergic neurons containing neuromelanin in the substantia nigra than unpigmented neurons are lost [68]. Impairment of the iron-chelating function of neuromelanin plays an essential role in Parkinson’s disease development, in which the amount of toxic iron bound to neuromelanin increases, thus rendering pigmented neurons susceptible to oxidative damage [18, 67, 74]. 2.2.2.5 Melanin in Adipose Tissue Eumelanin has been found to be synthesized in the visceral adipose tissue of morbidly obese patients [75]. However, the presence of pheomelanin was not investigated. The level of melanin biosynthesis in these tissues is three times higher in obese than in nonobese subjects. The expression of melanogenesisrelated genes (Tyr, Tyrp1, Dct, Rab27a, MC1R, and Melan-A) is also stronger in obese than in nonobese subjects. In adipose tissues, melanogenesis occurs in the cytosol rather than in melanosomes. The high levels of the melanogenic peptide α-MSH in the serum of obese subjects may account for the activation of the melanogenic pathway, because human adipocytes express the melanocortin receptor MC1R, which triggers α-MSHinduced melanogenesis in melanocytes. The ectopic synthesis of melanin in obese patients may be a compensatory mechanism conferring beneﬁts due to the antiinﬂammatory and oxidative damage-absorbing properties of melanin. The excess ROS due to obesity and the higher levels of cellular fat deposition may be neutralized by the melanin produced. According to the deﬁnition of melanocytes these cells may be considered nonclassical melanocytes.

2.3 Embryonic Development of Melanogenic Cells

In vertebrates, pigment cells have two embryonic origins: the neural crest and neuroepithelium. The pigment cells populating the retina are derived from the neuroepithelium. All other pigment cells are derived from a transient and pluripotent population of cells – the NCCs – considered to be the fourth germ layer. The neural crest delaminates from the roof of the developing neural tube and overlying ectoderm early in development along the rostrocaudal axis at the cephalic (from mid-diencephalon to rhombomere 8), vagal (from somite 1 to 7), truncal (from somite 8 to 28), and lumbo-sacral (posterior to somite 28) levels, to generate various lineages, such as neurons, glia, endocrine cells, bones, and melanocytes. Unlike other NCCs, melanocyte precursors can differentiate from the neural crest

37

38

2 Classical and Nonclassical Melanocytes in Vertebrates

irrespective of the region along the rostrocaudal axis in which they are located. The pigment cells derived from neural crest may give rise to classical and nonclassical melanocytes. 2.3.1 Classical Melanocytes 2.3.1.1

Early Determined Melanoblasts: The Dorsolateral Pathway

2.3.1.1.1 General Description Cutaneous melanocytes are derived from NCCs. In the truncal region, NCC precursors are found at the border between the neural ectoderm (or neural plate) and the non-neural ectoderm, in a region constituting the apex of the neural groove. In the trunk, the NCCs emerge when the dorsal end of the neural tube is fused, in the region corresponding to somites 8–28. This emergence follows an epithelialto-mesenchyme transition. The NCCs then enter a region called the migration staging area (MSA) located between the dorsal part of the somite, the lateral part of the neural tube, and the ventral part of the ectoderm [76]. In this region, the NCCs receive signals directing their migration and speciﬁcation. They take one of two migration pathways: the dorsoventral pathway, between the neural tube and the somites, and the dorsolateral pathway, between the ectoderm and the somites. Most NCCs migrate along the dorsoventral pathway to generate various types of cells, depending on the site of emergence along the rostrocaudal axis: sensory and sympathetic neurons, Schwann cells, endoneural ﬁbroblasts, chromafﬁn cells of the medullo-surrenal gland, and smooth muscle cells. Cells migrating along the dorsolateral pathway give rise only to melanocytes [77]. The NCCs following the dorsolateral pathway delaminate later than those following the dorsoventral pathway. Some of the NCCs produce Wnt1, Pax3, and Sox10. These cells give rise to the limited number of founder melanoblasts, which develop at about E8.5 [78]. Melanoblasts are nonpigmented cells committed to become melanocytes in the absence of external events preventing this determination. Founder melanoblasts begin to proliferate, generating precursor melanoblasts in the MSA. At E10, these melanoblasts produce Kit and Mitf. At around E10.5, they express a melanogenic enzyme marker, Dct. They proliferate in the MSA for about 1 day and then begin to migrate, colonizing the whole embryo. They migrate between the surface ectoderm and the somites (E10.5–E11), following a temporal rostrocaudal gradient. From E11.5, the migrating melanoblasts start entering the epidermis. They then cross the basal membrane and penetrate into the epidermis, from E12.5 in the mouse [79, 80]. From E15, a subset of melanoblasts migrates toward the matrix of the nascent hair follicles, where these cells express genes encoding proteins acting downstream from Mitf, such as Tyr and Tyrp1. Some follicular melanoblasts concentrate in the niche of the hair follicle, the “bulge.” These cells form the melanocyte stem cells and are responsible for maintaining homeostasis. Others migrate into the bulb of the hair follicle, where they differentiate into mature melanocytes, producing melanin in melanosomes at birth.

2.3 Embryonic Development of Melanogenic Cells

2.3.1.1.2 Genes Involved in Classical Melanocyte Development Many signaling molecules are required at all stages melanocyte development. They instruct the NCC to acquire a melanogenic fate and migrate along the dorsolateral pathway. They are necessary for melanocyte proliferation, differentiation, and survival. The following description is not exhaustive. Wnts and bone morphogenetic proteins (BMPs) are produced early in the development of the dorsal neural tube. BMPs favor the neuronal/glial differentiation of NCCs and inhibit pigment cell speciﬁcation, whereas Wnts play a dual role in promoting both sensory neurons and melanocytes. Melanoblasts arise from NCCs that delaminate late, at a time at which BMP4 is downregulated and Wnt signaling is maintained. Thus, temporal changes in the signals directing migration trajectories and cell fates in the dorsal neural tube may be involved in speciﬁcation of the melanocyte lineage [81]. Wnt/β-catenin signaling plays a major role in the determination of NCC cells to differentiate into melanocytes. Wnt1 and Wnt3a are expressed in the dorsal portion of the neural tube. The culture of mouse or avian NCCs with Wnt results in an increase in the number of pigment cells at the expense of other cell types [81]. In zebraﬁsh, the overproduction of β-catenin in migrating NCCs promotes the formation of pigment cells at the expense of neurons and glia [82]. Wnt signaling promotes pigment cell fate through direct activation of the melanocyte transcription factor Mitf-M by β-catenin (reviewed in [83]). Pax3, which is also produced early in the dorsal neural tube and in NCCs, is important for expansion of the melanoblast population [84] and for subsequent differentiation, as it transactivates Tyrp1 and acts in synergy with Sox10 to activate Mitf [85]. The FoxD3 transcription factor plays a critical role in determining the balance of NCC migration between the dorsolateral and dorsoventral migratory pathways. This factor inhibits melanoblast development. It is produced in the dorsal neural tube and in NCCs migrating early in the dorsoventral pathway. It continues to be produced in immature NC-derived cells, including Schwann cell precursors (SCPs), in which it may prevent differentiation. Its depletion from avian neural tubes promotes pigment cell development, whereas its overproduction prevents migration along the dorsolateral pathway [86]. Thomas and Erickson suggested that FoxD3 regulates the assignment of cells to a particular lineage by repressing Mitf gene expression in the early stages of neural crest development [87]. The two major signaling pathways stimulating the differentiation of melanocytes from the neural crest are the stem cell factor (SCF/Kitl)/Kit and Edn3/Ednrb pathways. Both Ednrb and Kit are involved in multiple steps in pigment cell development. Kit signaling is important at several timepoints in melanocyte development, having independent effects on both migration and survival [88–90]. Its role has been widely studied in the many Kit mutant mice generated to date, all of which present pigmentation defects. Kit is produced, together with Mitf, in both premigratory and migrating melanocytes, throughout their development [91], and its ligand, Kitl, is produced in a complementary manner in the dermamyotome, dermis, and hair follicles. Kit also seems to promote melanocyte

39

40

2 Classical and Nonclassical Melanocytes in Vertebrates

differentiation through post-translational modiﬁcations of Mitf [92]. Edn3 signaling is also important during many steps of melanocyte development, for migration, proliferation, survival, and differentiation [93]. In avian embryos, premigratory NCCs produce Ednrb, whereas Ednrb2 is activated in migrating and differentiated melanocytes [94]. In mice, Ednrb is required between E10 and E12.5, when melanoblasts migrate and proliferate [95]. Edn3/Ednrb also promotes Tyr gene expression [96]. Edn3 has a tissue distribution complementary to that of Ednrb, particularly in the epidermis [97, 98]. Edn3 is required earlier than Kit for the early migration of melanoblasts in the dermis [99]. These two signals are thought to inﬂuence each other. The gene encoding Mitf-M is considered to be the pigment cell master regulatory gene. Mitf-M directly regulates the expression of the melanogenic genes encoding Pmel17, Melan-A, and Trpm1, and the melanin synthesis enzyme genes encoding Tyr, Tyrp1, and Dct. Many Mitf mutant mice have been generated and all present pigmentation defects [100]. Mitf homologs are found in zebraﬁsh and Xenopus, and are also required for pigment cell development. In addition to regulating melanocyte differentiation, Mitf also regulates the survival of these cells, by controlling Bcl2 levels [101], and proliferation, by adjusting Ink4a and p21 levels [102, 103]. Many genes regulate Mitf expression and, thus, melanocyte development, either directly or indirectly. These genes include members of the Snail family (Snail and Slug), the Sox9, Sox10, and Ap2a genes. Sox10 transactivates Mitf [104], with which it acts in synergy to activate Dct [105]. A ﬁrst wave of differentiation of melanocyte occurs at birth. Other waves of differentiation occur periodically during life, melanocyte production is required to repopulate the adult hair follicles at each hair cycle. These melanocytes are produced from Dct-positive melanocyte stem cells present in the bulge area of the hair follicle [106]. Cell division and differentiation must be tightly regulated, to maintain the stem cell population and to guide their differentiation into melanocytes at each hair cycle. Stem cell maintenance is regulated by Mitf, Pax3, and Wnt signaling, all of which are also required for melanocyte development. Defects in the maintenance of this stem cell population may be responsible for hair graying [100, 107]. 2.3.1.2 Late Determined Melanoblasts: A Common Origin with SCPs and the Dorsoventral Migratory Pathway There seems to be a second pathway for the generation of melanocytes in the skin [108]. Growing nerves projecting throughout the body serve as a niche for stem/ progenitor cells, containing SCPs, which give rise to large numbers of skin melanocytes. These melanocytes develop after those following the dorsolateral pathway. They colonize the dorsal and lateral body walls, and seem to be the major melanocytes present in the limbs, as blocking the dorsolateral pathway has no effect on the number of melanoblasts in the limb bud. NCCs that delaminate early follow the ventral pathway of migration, with temporal distinctions between the various cells types generated in a ventral to dorsal direction, with sympathetic

2.3 Embryonic Development of Melanogenic Cells

neurons produced ﬁrst, followed by sensory neurons, while SCPs are produced throughout this period. SCPs are deﬁned as Sox10+ NCC-derived cells that are tightly associated with neuronal projections early in embryonic development and able to migrate long distances along the nerves. Sox10 is also expressed by multipotent NCCs and melanocytes. Melanocytes produced from the nerves innervating the skin originate from Sox10+/Krox20− SCPs of the nerve and produce Mitf and Sox10. They appear in the distal ventral ramus of the spinal nerve and then along the dorsal ramus nerves, which innervate the skin and axial muscles of the dorsal and lateral trunk. SCPs are generated due to a lack of neuronal speciﬁcation by Hmx1 gene function. This homeobox transcription factor may control a critical transcriptional switch between neuronal and glia melanocyte fates in the ventral NCC pathway. It is produced only in cells committed to a neuronal fate and is required for neurogenesis. Its absence leads to a shift in the balance between glial cells and neurons, increasing the numbers of SCPs and, thus, of melanoblasts. The determination of cell fate (SCP versus melanocyte) depends on contact with the nerve. For cells to be maintained as SCPs, they must be in contact with nerves. In the absence of signals provided by the nerve, some SCPs instead acquire a melanocyte fate. Promyelinating and myelinating Krox20+ Schwann cells do not normally differentiate into melanocytes, but they retain the competence to do so if nerve contact is lost. Krox20 expression deﬁnes a restriction of cell fate that normally prevents SCPs from differentiating into melanocytes. Cell fate determination also depends on interactions with neuregulin-1, which is expressed in neuron axonal membranes, and the ErbB2/ErbB3 complex receptor expressed by Schwann cells. Neuregulin signaling promotes the survival and proliferation of SCPs, and determines whether they have a glial or melanocyte fate. In addition, insulin-like growth factor-1 and platelet-derived growth factor, soluble factors produced in Schwann cells, have opposite effects to neuregulin-1, promoting melanocyte differentiation, survival and expansion from SCPs. Thus, Schwann cells are the source of both glia and melanocytes. This may help to account for the combination of changes in skin pigmentation and neurological disorders observed in patients with neuroﬁbromatosis type 1. 2.3.2 Nonclassical Melanocytes

With the exception of the RPE cells, which are derived from the neuroepithelium, nonclassical melanocytes, like classical melanocytes, are derived from the neural crest. These melanocytes may follow other migration routes, such as the dorsoventral pathway between the neural tube and the somites in the vagal region. They also differ from classical melanocytes in terms of their signaling pathway dependence. In general, decreases in Kit receptor tyrosine kinase signaling affect melanocyte development. However, the melanocytes in the eye, ear, and Harderian gland seem

41

42

2 Classical and Nonclassical Melanocytes in Vertebrates

to be less sensitive to Kit signaling than cutaneous melanocytes, as their precursors produce very small amounts of Kit and they are stimulated more effectively by Edn3 or hepatocyte growth factor (HGF) signals. Inhibition of these pathways speciﬁcally prevents the growth and differentiation of these noncutaneous melanocytes. Conversely, the induction of Edn3 or HGF expression in mouse skin and epithelial tissues promotes the survival and differentiation of noncutaneous and dermal melanocytes, with no similar effect on epidermal melanocytes, regardless of Kit signaling. Dermal melanocytes behave more like those of noncutaneous tissues than like epidermal melanocytes. There seem to be two major melanocyte populations: Kit-sensitive cutaneous melanocytes in the epidermis, and weakly Kit-sensitive noncutaneous and dermal melanocytes. These molecular differences between noncutaneous and dermal melanocytes, on the one hand, and epidermal melanocytes, on the other, may be important in the pathogenesis of melanocyterelated diseases and melanomas [109]. Little is known about the embryonic development of nonclassical melanocytes in mice and humans. A little information is available concerning the origin of the melanocytes found in the eye and the heart, but almost nothing is known about the origin of the melanocytes found in the brain or inner ear. Even less is known about the origin of nonclassical melanocytes in other species. 2.3.2.1 Melanocytes of the Murine Eye Melanocytes found in the eye (RPE and uveal melanocytes) have two different origins: neuroepithelium and neural crest. 2.3.2.1.1 Melanocytes of the RPE RPE cells are generated directly from the optic neuroepithelium and remain embedded in their local epithelial environment. An evagination of the neuroepithelium in the ventral forebrain is at the origin of the vertebrate eye. The optic neuroepithelium will be partitioned into three distinct territories, soon after the formation of the evagination: the distal territory will give rise to the future retina, the proximal territory to the optic stalk, and the dorsal territory to the RPE. The transition zone between the future retina and the RPE, the ciliary margin zone (CMZ), is at the origin of the ciliary body and the iris [110]. Eye development in the mouse starts at around E7.5. The neuroepithelium evaginates laterally in a dorsal-distal direction and extends toward the surface ectoderm, generating the optic stalk (which will become the optic nerve) proximally and the optic vesicle (which will become neuroretina, RPE, ciliary body, and iris) distally. The distal-most part of the vesicle forms the optic cup once it comes into contact with the surface ectoderm. By E13.5, the outer wall of the cup consists entirely of RPE. At E9.5, Mitf, a key pigmentation gene essential for skin melanocyte development, is expressed throughout the budding optic vesicle, together with Pax2 and Pax6. However, as early as E10.5, its expression is downregulated in the future retina, but becomes stronger in the future RPE. By contrast, Pax2 and Pax6 are downregulated in the future RPE, with Pax2 becoming concentrated in the optic stalk and Pax6 in the future retina [111].

2.3 Embryonic Development of Melanogenic Cells

Genes Involved in RPE Development The patterning of the optic neuroepithelium results from exposure to various extracellular ligands produced either by the neuroepithelium itself or by the surrounding tissues. Indeed, the extraocular mesenchyme promotes RPE development by expressing activins [112]. Hedgehog (HH) proteins have been found in the RPE of zebraﬁsh, Xenopus, and mouse, and their receptors and signal transduction proteins are expressed in the corresponding areas [113]. HH signaling promotes RPE formation [114, 115]. Fibroblast growth factors (FGFs) are negative regulators of RPE development. They are expressed in the surface ectoderm overlying the eye primordium and the corresponding receptors are found in the future neuroretina. In addition, the future retina itself produces FGFs. Despite this retinal expression, however, removal of the surface ectoderm and, thus, of a major source of FGFs, leads to the conversion of the presumptive neuroretina into pigmented RPE-like cells in both chicken embryos and optic vesicles harvested from mouse embryos. Conversely, exposure of the presumptive RPE to FGF leads to its development into neuroretina [116]. Nevertheless, FGFs are highly redundant in the eye. As a result, genetic studies have shown that no single ligand/receptor pair seems to be critical. The optic vesicle initially displays a high degree of developmental plasticity, indicating that each cell in the neuroepithelium can, in principle, respond to each of these ligands. The cells are, thus, initially equipotent. This raises questions about how the RPE is separated from the neuroretina distally and from the optic stalk proximally. Mitf is a critical transcription factor in the eye, playing a key role in separating the neuroretina from the RPE. It is produced in large amounts in the RPE [117, 118] and mutations rendering this molecule nonfunctional result in a hyperproliferating, unpigmented RPE, the dorsal part of which assumes a neuroretinal fate. These RPE abnormalities lead to small eyes. For more details on Mitf regulation in the RPE, see [110]. Vax proteins are responsible for separating the optic stalk from the RPE. Vax1 is conﬁned to the optic stalk, whereas Vax2 is restricted to the ventral neuroretina [119, 120]. The expression patterns of the Vax genes are therefore complementary to that of the Mitf gene, which is retained in the RPE, with a sharp boundary toward the optic stalk. Mitf and Vax are involved in mutual repression, leading to the formation of sharp boundaries between the optic stalk and RPE. The Kit and Edn3 signaling pathways, which are essential for melanocyte development, do not seem to be important in the RPE. In the RPE, cell proliferation is compatible with pigmentation and, thus, with differentiation. Mitf, which is essential for RPE proliferation, is also involved in the regulation of pigment gene expression and pigmentation in the RPE [118]. 2.3.2.1.2 Uveal Melanocytes Uveal melanocytes are neural crest-derived cells that migrate towards the eye during development. In humans, uveal melanogenesis begins in the 20th week of embryonic development and ﬁnishes some weeks after delivery. This is the reason for which deﬁnitive eye color – iris color – cannot be determined until the age of 6 months.

43

44

2 Classical and Nonclassical Melanocytes in Vertebrates

Choroid The choroid accounts for the vast majority of uveal melanocytes. The RPE is essential in the development of the choroid. Molecular interactions between the RPE and periocular mesenchyme are essential for melanocyte differentiation and vascular development in the choroid. The melanocytes of the choroid are derived from the cranial NCCs. In the human embryo, the pigmentation of choroidal melanocytes occurs late in gestation and is complete at birth. In the mouse, the pigmentation of choroidal melanocytes begins soon after birth and is complete by two weeks of age. In the absence of the RPE – if FGF9 is expressed in the presumptive RPE, resulting in a switch to neural retina, for example – the choroid fails to develop, demonstrating the requirement for the RPE as a source of inductive signals for choroid development [49]. Iris The iris consists of ﬁve cell layers, two of which are pigmented: the stroma and the posterior pigment epithelium [48]. The iris pigment epithelium consists of a double layer of cuboidal pigmented cells that are tightly fused. These cells are of neuroectodermal origin, and are derived from the CMZ at the anterior end of the optic cup. By contrast, the stromal melanocytes are derived from NCCs and migrate through the uveal tract during development. Ciliary Body Like the iris, the ciliary body – a structure linking the iris to the choroid and forming part of the uveal tract – has a pigmented epithelium derived from the CMZ and stromal melanocytes derived from the neural crest. 2.3.2.2 Melanocytes of the Murine Heart The melanocytes of the heart are derived from cardiac NCCs, which leave the neural tube between the postotic area and the ﬁrst four somites (a domain that overlaps the cranial and vagal neural crest). Dct-positive melanoblasts have been found in the postotic area extending through the second and third branchial arches, and in a more posterior position, in the region of somites 2 and 3. They have also been seen migrating close to the sixth branchial arch arteries. At E11.5, they are found close to the endocardial cushions and appear to enter the heart via the septum primum. These cardiac melanoblasts seem to emigrate from the neural tube earlier than those migrating dorsolaterally to colonize the skin. At E12.5, the cardiac melanoblasts are found in the AV endocardial cushions, and in the areas surrounding the aortic sac and common cardinal vein. They are also found on the dorsal side of the right atrium, apparently traveling along the anterior cardinal vein and the sixth branchial arch artery [61]. By E13.5, these cells are found in the pulmonary veins and septal leaﬂets of the AV valves. Between E14.5 and E16.5, they colonize the posterior atrium and atrial-pulmonary venous anastomoses and, by late gestation, they are found in regions of the compact AV node and sinoatrial node [62]. This pattern of expression is subsequently maintained throughout adulthood. Cardiac melanoblasts originate from the same axial levels as cardiac NCCs, with which they share migratory pathways, but they constitute a separate population in terms of their level of commitment, the timing of their arrival in the heart, and their dependence on signaling. They seem to be derived

2.4 Transfer of Melanin from Classical and Nonclassical Melanocytes

from a subpopulation of cardiac neural crest precursors that do not express Wnt1 and Pax3 at early timepoints [61, 62]. 2.3.2.3 Other Nonclassical Murine Melanocytes In the brain, the melanocytes in the sympathetic cephalic ganglia and leptomeninges are derived from cephalic NCCs. In the inner ear, the melanocytes of the stria vascularis are derived from cephalic NCCs. 2.3.2.4 Other Organisms Chromatophores are just one of a number of cell types generated by the neural crest during embryonic development in lower vertebrates. They migrate long distances to populate many organs of the body, including the skin, eye, ear, and brain. In ﬁsh and Xenopus, pigment is produced in the chromatophores before migration is completed. Chromatophores leave the neural crest in waves and follow either a dorsolateral route through the dermis, entering the ectoderm through small holes in the basal lamina, or a ventromedial route between the somites and the neural tube. In ﬁsh, the melanophores appear laterally at about 24 h of development and are subsequently found along the trunk. The xanthophores are the next to appear, followed by the iridophores [121]. The melanophores of the RPE of the eye constitute an exception. They are not derived from the neural crest and originate instead from the optic cup generated from the neuroepithelium, as in higher vertebrates. The development of chromatoblasts – the precursor cells of chromatophores – seems to be dependent on the same signaling pathways as that of higher vertebrate melanocytes, because Kit, Sox10, and Mitf play an important role in controlling chromatophore differentiation in ﬁsh [121, 122]. Defects in these proteins may result in chromatophores being entirely absent or absent from particular regions, resulting in a leukistic disorder.

2.4 Transfer of Melanin from Classical and Nonclassical Melanocytes

Melanin is actively transferred principally from epidermal melanocytes. We will not focus here on the biogenesis of melanosomes and the transport of the melanin, which are dealt with in other chapters of the book (see Chapters 9 and 10). Instead, we describe here the conditions and proteins essential for the correct transfer of melanin from the donor cells to the recipient cells. The presence or absence of crucial players may account for the lack of melanin transfer from nonclassical melanocytes. The epidermal melanin unit is a functional unit consisting of one melanocyte and several neighboring keratinocytes. The melanocytes synthesize the melanin and donate it to the recipient cells, the epidermal keratinocytes, which acquire and retain most of the melanin in the skin. The melanosome is a unique membrane-bound organelle in which melanin biosynthesis takes place. Melanin is a complex pigment that protects the skin

45

46

2 Classical and Nonclassical Melanocytes in Vertebrates

against the potentially harmful effects of light, by absorbing and scattering light and reducing the production of ROS [123]. For this to be achieved, melanosomes must be transported to the tips of melanocyte dendrites and transferred into the surrounding keratinocytes [124]. This transfer is essential for photoprotection of the skin and the maintenance of normal skin color. Melanosomes are lysosomerelated organelles [125] and they display four stages of maturation, based on their appearance under an electron microscope and their melanization status [1]. Stage I melanosomes (or premelanosomes) are round vesicles found in the perinuclear zone, where they probably develop from the endoplasmic reticulum [2], or from endosomes [3]. The transition of premelanosomes to stage II of maturation involves an elongation of the vesicles and the organization of a distinct ﬁbrillar matrix, but no melanogenesis activity is detected, despite the presence of Tyr. Melanin is ﬁrst synthesized in stage III, when it is deposited on the ﬁbrillar matrix. In stage IV, deposition continues until the internal structure is no longer visible [4, 5]. 2.4.1 Melanosome Transport

Skin pigmentation is dependent on the transport of melanosomes within melanocytes, from the area of synthesis close to the center of the cell to the peripheral dendrites, before transfer to keratinocytes [126–129]. The intracellular transport of melanosomes is directed by two major polarized cytoskeletal macromolecular polymers – actin ﬁlaments and microtubules – composed of actin monomers and dimers of α- and β-tubulin, respectively. Melanosomes are shuttled along the microtubules by certain members of two major families of motor proteins: dyneins and kinesins [130]. Transport along actin ﬁlaments is mediated by myosins [131]. Dynein and kinesin act as short cross-bridge structures connecting the organelle to the microtubules [130, 132]. The net transport of melanosomes depends on the balance between the forces directed toward the center and those directed toward the periphery in melanosomes. Myosin Va is an important motor in the actin ﬁlament-based transport of organelles [133]. It captures melanosomes delivered to the periphery via the centrifugal component of microtubule-dependent movement, by mediating their interaction with actin ﬁlaments forming a cortical shell running the length of the dendrites right to their tips [134]. The long-range, bidirectional, microtubule-dependent movements of melanosomes, coupled with the actomyosin Va-dependent capture of melanosomes in the periphery, is the principal mechanism underlying the centrifugal transport and peripheral accumulation of melanosomes in melanocytes. 2.4.2 Melanosome Transfer

In mammals, skin and coat colors are regulated by melanosome transfer from melanocytes to neighboring keratinocytes and/or to the hair bulb cells. In humans,

2.4 Transfer of Melanin from Classical and Nonclassical Melanocytes

pigment is transferred principally to protect against the damaging effects of UV radiation from the sun [135]. Melanocytes close to the basal lamina in the skin export melanin to the surrounding keratinocytes in the epidermis, resulting in a tanning effect. The melanized keratinocytes are constantly shed and renewed, thus creating a need to produce new melanin continually. The number of melanocytes is similar in humans with different skin types, the differences between ethnic groups resulting mostly from differences in the melanogenic activity of each melanocyte and the size and maturation of the melanosomes [135, 136]. Most studies on pigment transfer have been carried out in mammals – principally mice and humans – but some investigations have focused on amphibians [41, 137]. Several models of melanosome transfer are based on the assumption that melanosomes can be correctly retained close to the cell membrane. This maintenance of melanosomes at the cell membrane is achieved by the association of melanosomes with actin ﬁlaments, which are abundant at the tips of the dendrites. 2.4.2.1

Melanosome Transfer from Classical Melanocytes

2.4.2.1.1 Mechanisms of Melanin Transfer Melanocytes have many dendrites, making it possible for each melanocyte to maintain contact with several keratinocytes simultaneously. This allows for the phagocytosis of dendrite tips by the keratinocytes, but also makes it possible for large numbers of melanosomes to be maintained close to the cell membrane, facilitating other modes of transfer. The mechanism underlying melanosome transfer is less well understood than the mechanism of melanosome transport from the perinuclear area to the tips of the dendrites. Several models of the transfer of mature melanosomes to neighboring keratinocytes have been proposed, based on different model systems and observation methods: exocytosis, cytophagocytosis, fusion of plasma membranes, and transfer by membrane vesicles. The large number of models proposed reﬂects the possibility of several of these mechanisms operating simultaneously in the same organism or of different mechanisms operating in animals of different species or ages or in different tissues. The mechanisms of melanosome transfer are described in Chapter 10. 2.4.2.1.2 Molecular Players in Melanin Transfer The molecular and cellular mechanisms involved in melanosome transfer have been only partly elucidated, but membrane fusion between the melanosome and the plasma membrane of the melanocyte dendrite or the keratinocyte membrane seems likely to be involved in the transfer process. Melanogenic paracrine and autocrine cytokines have been discovered in vitro, mediating interactions between melanocytes and other types of skin cells. α-MSH, ACTH, basic FGF, nerve growth factor (NGF), endothelins, granulocyte-macrophage colony-stimulating factor (GM-CSF), SCF, leukemia inhibitory factor (LIF), and HGF have also been reported to be produced by keratinocytes and involved in regulating the proliferation and/ or differentiation of mammalian epidermal melanocytes [138]. Indeed, keratinocytes may produce and release many factors involved in regulating the proliferation

47

48

2 Classical and Nonclassical Melanocytes in Vertebrates

and differentiation of mammalian epidermal melanocytes through receptormediated signaling pathways. Foxn1 For many years, little was known about the role of epithelial cells in the development of pigmentary interactions. Until recently, it remained unclear whether epithelial cells were “passive,” acquiring pigment only if spontaneously provided by a melanocyte, or “active,” recruiting melanocytes and inducing the transfer of pigment. The group of Janice Brissette has reconsidered this issue of the passive/active role of keratinocytes in melanin transfer from melanocytes. They have shown that Foxn1, a transcription factor, is responsible for identifying pigment recipient cells and recruiting pigment donors to generate pigmentary units. Foxn1 stimulates keratinocytes to emit signals, one of which is FGF2. Melanocytes make use of these signals to recognize the Foxn1-positive cells as targets, to which they connect, via dendrites, and transfer pigment [139]. Foxn1 is the ﬁrst gene to be implicated in the uptake of pigment, but it is not the only one. Foxn1 is not universally expressed in the pigment recipient cells, being limited to only some of the keratinocytes. Other mechanisms must therefore be involved in the reception of pigment. PAR-2 It has been reported that keratinocytes express a proteinase-activated receptor 2 (PAR-2) [140, 141], from the large family of seven-transmembraneregion receptors that bind guanosine nucleotide-binding proteins. PAR-2 is widespread throughout the body, being found in the gastrointestinal tract, pancreas, kidneys, liver, lung, cardiovascular system, ovary, eye and brain [142–144]. The physiological activation of PAR-2 is induced by serine protease cleavage of its extracellular region, resulting in a new N-terminus that acts as a tethered ligand, free to interact with another region of the receptor, leading to activation. PAR-2 stimulation has been shown to induce a rearrangement of cytoskeletal proteins [145] and an increase in the phagocytic activity of keratinocytes. This increases melanin transfer, thereby increasing the overall pigmentation of the epidermis [145–147]. Intimate contact between melanocytes and keratinocytes is required for PAR-2-mediated pigmentation effects. PAR-2 is expressed in melanocytes only during interactions with keratinocytes [147]. Consistent with the role of PAR-2 in the phagocytosis of melanosomes by keratinocytes and with the stimulation of melanosome transfer by UV irradiation, UV upregulates PAR-2 production and induces its activity, both in vitro and in vivo. Proteases with PAR-2 cleavage activity are also induced in UV-treated keratinocytes [148]. The effect of UV on PAR-2 synthesis and activity is more signiﬁcant in coloredskinned individuals (skin phototype II and III) than in those with fair skin (skin phototype I) [148]. In vitro experiments have shown that PAR-2 activation stimulates serine protease secretion by keratinocytes, thus creating a positive feedback loop [148]. Analyses of the cellular signaling pathways regulating the effects mediated by PAR-2 have shown that PAR-2-mediated phagocytosis is Rho-dependent, Rho being a downstream mediator of PAR-2. PAR-2 activation results in a rapid and

2.4 Transfer of Melanin from Classical and Nonclassical Melanocytes

concentration-dependent increase in cAMP levels, but PAR-2-mediated Rho activation is not linked to the protein kinase A signaling pathway [149]. In addition to inducing phagocytosis, PAR-2 mediates skin pigmentation by stimulating melanocyte dendrite development through the release of prostaglandins – principally prostaglandin E2 (PGE2) and PGE2α [150], which are released by keratinocytes in response to UV stimulation [151, 152]. KGF The inhibition of serine protease activity does not completely prevent melanosome transfer [146], suggesting that PAR-2 is just one of a number of participants in the melanin transfer process. Indeed, keratinocyte growth factor (KGF/FGF7) has been implicated in melanosome transfer, in which it exerts its effects principally by acting on the recipient keratinocytes [153]. KGF is secreted by dermal ﬁbroblasts [154, 155] and acts by binding to the KGF receptor (KGFR), a splicing variant of ﬁbroblast growth factor receptor 2. KGF binding, together with UV exposure, triggers the activation and internalization of KGFR, thereby inducing phagocytosis and melanosome transfer in vitro [153]. The phagocytosis promoted by KGF involves actin reorganization and the activation of both Rho- and Cdc42/Rac-mediated mechanisms [153]. Furthermore, KGFR is detected in perinuclear phagosomes, suggesting a possible role in the intracellular translocation of melanosomes from the periphery to the central area of the keratinocytes, to protect the nucleus [156]. KGFR activity and signaling are observed in keratinocytes from both light and dark skins, but expression levels are higher in light than in dark skins. This accounts for the weaker lower level of responsiveness of dark keratinocytes to KGF treatment and further demonstrates that the receptor is responsible for differences in melanosome uptake, but that its presence on the membrane is not sufﬁcient in itself [157]. The stronger effect of KGF on melanosome transfer to keratinocytes from light skin than on that to keratinocytes from dark skin suggests that other pathways, in addition to the well known PAR-2-activated pathway [148], must be activated in light skin, thereby enhancing protection against UV-induced damage. Cadherins Cadherins are calcium-dependent transmembrane glycoproteins involved in promoting cell–cell adhesion via homophilic binding and acting as the transmembrane components of cell–cell junctions [158]. Human keratinocytes [159] and melanocytes have been shown to express both E-cadherin and P-cadherin, and these proteins have been shown to play a major role in the establishment of melanocyte–keratinocyte adhesion. P-cadherin seems to be less important than E-cadherin, which has been shown to play a major role in the adhesion of melanocytes to keratinocytes [160]. The role of E-cadherin in melanosome transfer is highlighted by the dissociation of melanocytes and keratinocytes in the acantholytic lesions of Darier’s disease [160, 161], in which there is a loss of keratinocyte– melanocyte adhesion that results in a loss of pigment transfer. Lectins and Glycoproteins In addition to cadherins, the melanocyte–keratinocyte adhesion binds lectins and glycoproteins, via sugar-binding adhesion receptors

49

50

2 Classical and Nonclassical Melanocytes in Vertebrates

on the surface of the delimiting membranes. Plasma membrane lectins and neoglycoproteins have been shown to be involved in melanocyte–keratinocyte recognition and to facilitate melanin transfer [162]. Some of these membrane receptors have been shown to be upregulated in melanoma cells in response to UV irradiation [163], which is known to increase melanosome transfer [164]. The membrane lectins binding fucose neo glycoproteins are strongly expressed on melanoma cells and only weakly expressed on keratinocytes, consistent with the expression of fucose receptors on melanocytes and the attachment of fucose residues to proteins on the keratinocytes [163]. The addition of speciﬁc lectins and neoglycoproteins to cocultures of melanocytes and keratinocytes may disrupt melanosome transfer. These ligands seem to bind their speciﬁc plasma membrane receptors and interfere with the melanocyte and keratinocyte intercellular contacts essential for melanosome transfer [162]. This inhibition is reversible and may be enhanced by adding niacinamide [165] SNAREs and Rab Membrane fusion events and exocytosis are often mediated by soluble N-ethylmaleimide-sensitive factor attachment protein receptors (SNAREs) and Rab GTPases [166]. These proteins are present in melanocytes, suggesting that they may regulate the targeting of melanosomes to the plasma membrane and the transfer of melanosomes to keratinocytes [148]. SNAREs are mostly membrane-anchored proteins that serve as membrane receptors. On the basis of their distribution, SNARE proteins are classiﬁed as either vesicle SNAREs (VAMP/ synaptobrevin) or target SNAREs (syntaxins and SNAP proteins). VAMP proteins can bind to syntaxins and SNAP proteins, forming core complexes that regulate membrane fusion [167]. Melanocytes have been shown to produce both vesicle SNARE (VAMP-2) and target SNARE (SNAP-25, SNAP-23, and syntaxin-4) proteins [148, 168]. The treatment of melanocytes with α-MSH leads to an increase in the production of several SNARE proteins, consistent with a role for these proteins in melanosome transfer [169]. VAMP-2, on melanosomes, and SNAP-23, on the plasma membrane, associate in melanocytes to facilitate membrane fusion. Despite its presence on the plasma membrane of melanocytes, syntaxin-4 does not associate with VAMP-2 and SNAP-23, suggesting that another, as yet unidentiﬁed, syntaxin may be involved. Following fusion, melanosomes may be released into the extracellular space, to be taken up by keratinocytes through phagocytosis or a membrane fusion event. Rab GTPases constitute another family of proteins involved in membrane fusion, particularly in the tethering and attachment of melanosomes to the plasma membrane, before fusion actually occurs. Rab3a inhibits regulated exocytosis [170–172]. Its expression is downregulated by UV irradiation, inhibiting melanosome transport and transfer [148, 173]. Rab GTPases involved in melanosome transport include Rab27a, which mediates myosin Va binding to melanosomes through another linker protein, melanophilin. Rab27a keeps the melanosomes attached to the plasma membrane, which is particularly important for exocytosis.

2.4 Transfer of Melanin from Classical and Nonclassical Melanocytes

2.4.2.1.3 Melanosome Positioning within Keratinocytes Following melanosome uptake by keratinocytes, there may be one melanosome per phagosome or several melanosomes within a single large phagosome [174]. Melanosome sorting seems to differ between skin types, as dark-skinned individuals store their large melanosomes individually in single vesicles, whereas fairskinned individuals store smaller melanosomes in larger vesicles within keratinocytes [136, 175]. Within the keratinocytes, melanosomes aggregates towards the apical pole of the nucleus, forming a supranuclear cap protecting the cell from damage due to UV irradiation. They are degraded when the keratinocytes undergo terminal differentiation and desquamation. 2.4.2.2 Transfer of Melanin from Nonclassical Melanocytes Melanin and melanocytes are not restricted to the skin and hair (classical melanocytes) as previously mentioned. They are also found in the eyes (uveal tract, choroid, ciliary body and iris), inner ear (stria vascularis and modulus of the cochlea, dark cells of the vestibular organ), leptomeninges of the entire brain, heart, and adipose tissue (nonclassical melanocytes). Melanocytes are known, as described above, to produce melanin, which they transfer to keratinocytes to protect these cells from the harmful effects of sun exposure [176]. The presence of melanocytes in organs that are not exposed to the sun suggests that these cells must have other functions [35]. Indeed, whereas melanin is continually produced and secreted in the skin and hair, the melanosomes of nonclassical melanocytes are not transferred to the surrounding cells. They are instead retained in the melanocytes, accumulating in the cytoplasm of these cells within the various stromas [63, 67]. The absence of melanin transfer from nonclassical melanocytes to the neighboring cells may be accounted for by the lack of keratinocytes continually producing speciﬁc factors modulating melanocyte–keratinocyte contact and controlling the mechanism of melanin transfer to keratinocytes. For example, the cells surrounding melanocytes in the heart are not pigmented, so melanin does not seem to be transferred to or to accumulate in the neighboring cells in this organ, although electron microscopy experiments would be required to conﬁrm this. It has been shown that the skin cells surrounding melanocytes produce speciﬁc proteins, such as Foxn1 [139]. Reverse transcription polymerase chain reaction experiments failed to detect Foxn1 mRNA in the heart [63], potentially accounting for the lack of melanin transfer from melanocytes to adjacent cells in the heart. Furthermore, the PAR-2 gene, which is widely expressed in the human body, is only very weakly expressed in the heart and its expression has never been detected in the brain. PAR-2 expression in melanocytes has also been reported to be induced only on interaction with keratinocytes [147]. In conclusion, melanocytes alone, in the absence of keratinocytes, have limited pigmentation power. They require the effective modulators of melanocyte– keratinocyte interactions and melanin transfer produced by the neighboring keratinocytes.

51

52

2 Classical and Nonclassical Melanocytes in Vertebrates

References 1 Kushimoto, T., Basrur, V., Valencia, J.,

2

3

4

5

6

7

8

9

Matsunaga, J., Vieira, W.D., Ferrans, V.J., Muller, J., Appella, E., and Hearing, V.J. (2001) A model for melanosome biogenesis based on the puriﬁcation and analysis of early melanosomes. Proc. Natl. Acad. Sci. USA, 98, 10698–10703. Slominski, A., Tobin, D.J., Shibahara, S., and Wortsman, J. (2004) Melanin pigmentation in mammalian skin and its hormonal regulation. Physiol. Rev., 84, 1155–1228. Raposo, G., Tenza, D., Murphy, D.M., Berson, J.F., and Marks, M.S. (2001) Distinct protein sorting and localization to premelanosomes, melanosomes, and lysosomes in pigmented melanocytic cells. J. Cell Biol., 152, 809–824. Setaluri, V. (2003) The melanosome: dark pigment granule shines bright light on vesicle biogenesis and more. J. Invest. Dermatol., 121, 650–660. Hearing, V.J. (2005) Biogenesis of pigment granules: a sensitive way to regulate melanocyte function. J. Dermatol. Sci., 37, 3–14. Schmitz, S., Thomas, P.D., Allen, T.M., Poznansky, M.J., and Jimbow, K. (1995) Dual role of melanins and melanin precursors as photoprotective and phototoxic agents: inhibition of ultraviolet radiation-induced lipid peroxidation. Photochem. Photobiol., 61, 650–655. Salinas, C., Garcia-Borron, J.C., Solano, F., and Lozano, J.A. (1994) Dopachrome tautomerase decreases the binding of indolic melanogenesis intermediates to proteins. Biochim. Biophys. Acta, 1204, 53–60. Bush, W.D. and Simon, J.D. (2007) Quantiﬁcation of Ca2+ binding to melanin supports the hypothesis that melanosomes serve a functional role in regulating calcium homeostasis. Pigment Cell Res., 20, 134–139. Gilchrest, B.A., Eller, M.S., Geller, A.C., and Yaar, M. (1999) The pathogenesis of melanoma induced by ultraviolet radiation. N. Engl. J. Med., 340, 1341–1348.

10 Landi, M.T., Baccarelli, A., Tarone, R.E.,

11

12

13

14

15

16

17

18

19

Pesatori, A., Tucker, M.A., Hedayati, M., and Grossman, L. (2002) DNA repair, dysplastic nevi, and sunlight sensitivity in the development of cutaneous malignant melanoma. J. Natl. Cancer Inst., 94, 94–101. Robinson, S.J. and Healy, E. (2002) Human melanocortin 1 receptor (MC1R) gene variants alter melanoma cell growth and adhesion to extracellular matrix. Oncogene, 21, 8037–8046. Yamaguchi, Y., Brenner, M., and Hearing, V.J. (2007) The regulation of skin pigmentation. J. Biol. Chem., 282, 27557–27561. Agar, N. and Young, A.R. (2005) Melanogenesis: a photoprotective response to DNA damage? Mutat. Res., 571, 121–132. Eller, M.S., Ostrom, K., and Gilchrest, B.A. (1996) DNA damage enhances melanogenesis. Proc. Natl. Acad. Sci. USA, 93, 1087–1092. Cui, R., Widlund, H.R., Feige, E., Lin, J.Y., Wilensky, D.L., Igras, V.E., D’Orazio, J., Fung, C.Y., Schanbacher, C.F., Granter, S.R., and Fisher, D.E. (2007) Central role of p53 in the suntan response and pathologic hyperpigmentation. Cell, 128, 853–864. Tolleson, W.H. (2005) Human melanocyte biology, toxicology, and pathology. J. Environ. Sci. Health C, 23, 105–161. Ernfors, P. (2010) Cellular origin and developmental mechanisms during the formation of skin melanocytes. Exp. Cell Res., 316, 1397–1407. Plonka, P.M., Passeron, T., Brenner, M., Tobin, D.J., Shibahara, S., Thomas, A., Slominski, A., Kadekaro, A.L., Hershkovitz, D., Peters, E., Nordlund, J.J., Abdel-Malek, Z., Takeda, K., Paus, R., Ortonne, J.P., Hearing, V.J., and Schallreuter, K.U. (2009) What are melanocytes really doing all day long…? Exp. Dermatol., 18, 799–819. Kobayashi, N., Nakagawa, A., Muramatsu, T., Yamashina, Y., Shirai, T., Hashimoto, M.W., Ishigaki, Y.,

References

20

21

22

23

24

25

26

27

Ohnishi, T., and Mori, T. (1998) Supranuclear melanin caps reduce ultraviolet induced DNA photoproducts in human epidermis. J. Invest. Dermatol., 110, 806–810. Bustamante, J., Bredeston, L., Malanga, G., and Mordoh, J. (1993) Role of melanin as a scavenger of active oxygen species. Pigment Cell Res., 6, 348–353. Millington, G.W. (2006) Proopiomelanocortin (POMC): the cutaneous roles of its melanocortin products and receptors. Clin. Exp. Dermatol., 31, 407–412. Suzuki, I., Tada, A., Ollmann, M.M., Barsh, G.S., Im, S., Lamoreux, M.L., Hearing, V.J., Nordlund, J.J., and Abdel-Malek, Z.A. (1997) Agouti signaling protein inhibits melanogenesis and the response of human melanocytes to alpha-melanotropin. J. Invest. Dermatol., 108, 838–842. Kadekaro, A.L., Kavanagh, R., Kanto, H., Terzieva, S., Hauser, J., Kobayashi, N., Schwemberger, S., Cornelius, J., Babcock, G., Shertzer, H.G., Scott, G., and Abdel-Malek, Z.A. (2005) Alphamelanocortin and endothelin-1 activate antiapoptotic pathways and reduce DNA damage in human melanocytes. Cancer Res., 65, 4292–4299. Bohm, M., Wolff, I., Scholzen, T.E., Robinson, S.J., Healy, E., Luger, T.A., Schwarz, T., and Schwarz, A. (2005) Alpha-melanocyte-stimulating hormone protects from ultraviolet radiationinduced apoptosis and DNA damage. J. Biol. Chem., 280, 5795–5802. Mackintosh, J.A. (2001) The antimicrobial properties of melanocytes, melanosomes and melanin and the evolution of black skin. J. Theor. Biol., 211, 101–113. Le Poole, I.C., Mutis, T., van den Wijngaard, R.M., Westerhof, W., Ottenhoff, T., de Vries, R.R., and Das, P.K. (1993) A novel, antigen-presenting function of melanocytes and its possible relationship to hypopigmentary disorders. J. Immunol., 151, 7284–7292. Aubock, J., Niederwieser, D., Romani, N., Fritsch, P., and Huber, C. (1985)

28

29

30

31

32

33

34

35

36

Human interferon-gamma induces expression of HLA-DR on keratinocytes and melanocytes. Arch. Dermatol. Res., 277, 270–275. Rheins, L.A. and Nordlund, J.J. (1986) Modulation of the population density of identiﬁable epidermal Langerhans cells associated with enhancement or suppression of cutaneous immune reactivity. J. Immunol., 136, 867–876. Slominski, A. and Wortsman, J. (2000) Neuroendocrinology of the skin. Endocr. Rev., 21, 457–487. Slominski, A., Wortsman, J., Luger, T., Paus, R., and Solomon, S. (2000) Corticotropin releasing hormone and proopiomelanocortin involvement in the cutaneous response to stress. Physiol. Rev., 80, 979–1020. Grando, S.A., Pittelkow, M.R., and Schallreuter, K.U. (2006) Adrenergic and cholinergic control in the biology of epidermis: physiological and clinical signiﬁcance. J. Invest. Dermatol., 126, 1948–1965. Slominski, A., Zbytek, B., Pisarchik, A., Slominski, R.M., Zmijewski, M.A., and Wortsman, J. (2006) CRH functions as a growth factor/cytokine in the skin. J. Cell Physiol., 206, 780–791. Takeda, K., Yokoyama, S., Aburatani, H., Masuda, T., Han, F., Yoshizawa, M., Yamaki, N., Yamamoto, H., Eguchi, N., Urade, Y., and Shibahara, S. (2006) Lipocalin-type prostaglandin D synthase as a melanocyte marker regulated by MITF. Biochem. Biophys. Res. Commun., 339, 1098–1106. Beuckmann, C.T., Aoyagi, M., Okazaki, I., Hiroike, T., Toh, H., Hayaishi, O., and Urade, Y. (1999) Binding of biliverdin, bilirubin, and thyroid hormones to lipocalin-type prostaglandin D synthase. Biochemistry, 38, 8006–8013. Takeda, K., Takahashi, N.H., and Shibahara, S. (2007) Neuroendocrine functions of melanocytes: beyond the skin-deep melanin maker. Tohoku J. Exp. Med., 211, 201–221. Buffey, J.A., Messenger, A.G., Taylor, M., Ashcroft, A.T., Westgate, G.E., and MacNeil, S. (1994) Extracellular matrix derived from hair and skin ﬁbroblasts stimulates human skin melanocyte

53

54

2 Classical and Nonclassical Melanocytes in Vertebrates

37

38

39

40

41

42

43

44

45

tyrosinase activity. Br. J. Dermatol., 131, 836–842. Archambault, M., Yaar, M., and Gilchrest, B.A. (1995) Keratinocytes and ﬁbroblasts in a human skin equivalent model enhance melanocyte survival and melanin synthesis after ultraviolet irradiation. J. Invest. Dermatol., 104, 859–867. Hedley, S.J., Layton, C., Heaton, M., Chakrabarty, K.H., Dawson, R.A., Gawkrodger, D.J., and MacNeil, S. (2002) Fibroblasts play a regulatory role in the control of pigmentation in reconstructed human skin from skin types I and II. Pigment Cell Res., 15, 49–56. Yamaguchi, Y., Passeron, T., Hoashi, T., Watabe, H., Rouzaud, F., Yasumoto, K., Hara, T., Tohyama, C., Katayama, I., Miki, T., and Hearing, V.J. (2008) Dickkopf 1 (DKK1) regulates skin pigmentation and thickness by affecting Wnt/beta-catenin signaling in keratinocytes. FASEB J., 22, 1009–1020. Yamaguchi, Y., Itami, S., Watabe, H., Yasumoto, K., Abdel-Malek, Z.A., Kubo, T., Rouzaud, F., Tanemura, A., Yoshikawa, K., and Hearing, V.J. (2004) Mesenchymal–epithelial interactions in the skin: increased expression of dickkopf1 by palmoplantar ﬁbroblasts inhibits melanocyte growth and differentiation. J. Cell Biol., 165, 275–285. Aspengren, S., Hedberg, D., and Wallin, M. (2006) Studies of pigment transfer between Xenopus laevis melanophores and ﬁbroblasts in vitro and in vivo. Pigment Cell Res., 19, 136–145. Yasutomi, M. (1987) Migration of epidermal melanophores to the dermis through the basement membrane during metamorphosis in the frog, Rana japonica. Pigment Cell Res., 1, 181–187. Chow, R.L. and Lang, R.A. (2001) Early eye development in vertebrates. Annu. Rev. Cell Dev. Biol., 17, 255–296. Martinez-Morales, J.R., Rodrigo, I., and Bovolenta, P. (2004) Eye development: a view from the retina pigmented epithelium. Bioessays, 26, 766–777. Bok, D. (1993) The retinal pigment epithelium: a versatile partner in vision. J. Cell Sci. Suppl., 17, 189–195.

46 Jeffery, G. (1997) The albino retina: an

47

48

49

50

51

52

53

54 55

56

57

abnormality that provides insight into normal retinal development. Trends Neurosci., 20, 165–169. Biesemeier, A., Kreppel, F., Kochanek, S., and Schraermeyer, U. (2010) The classical pathway of melanogenesis is not essential for melanin synthesis in the adult retinal pigment epithelium. Cell Tissue Res., 339, 551–560. Sturm, R.A. and Larsson, M. (2009) Genetics of human iris colour and patterns. Pigment Cell Melanoma Res., 22, 544–562. Mouriaux, F., Saule, S., Desjardins, L., and Mascarelli, F. (2005) Normal and malignant choroidal melanocytes: from cell to clinical approach. J. Fr. Ophtalmol., 28, 781–793. Payne, A.P. (1994) The harderian gland: a tercentennial review. J. Anat., 185, 1–49. Tachibana, M. (1999) Sound needs sound melanocytes to be heard. Pigment Cell Res., 12, 344–354. Keithley, E.M., Ryan, A.F., and Feldman, M.L. (1992) Cochlear degeneration in aged rats of four strains. Hear. Res., 59, 171–178. Benedito, E., Jimenez-Cervantes, C., Perez, D., Cubillana, J.D., Solano, F., Jimenez-Cervantes, J., Meyer zum Gottesberge, A.M., Lozano, J.A., and Garcia-Borron, J.C. (1997) Melanin formation in the inner ear is catalyzed by a new tyrosine hydroxylase kinetically and structurally different from tyrosinase. Biochim. Biophys. Acta, 1336, 59–72. Steel, K.P. (1995) Hair-cell regeneration: cure for deafness? Lancet, 346, 325–326. Price, E.R. and Fisher, D.E. (2001) Sensorineural deafness and pigmentation genes: melanocytes and the Mitf transcriptional network. Neuron, 30, 15–18. Gill, S.S. and Salt, A.N. (1997) Quantitative differences in endolymphatic calcium and endocochlear potential between pigmented and albino guinea pigs. Hear. Res., 113, 191–197. Meyer zum Gottesberge, A.M. (1988) Physiology and pathophysiology of inner

References

58

59

60

61

62

63

64

65

ear melanin. Pigment Cell Res., 1, 238–249. Uehara, S., Izumi, Y., Kubo, Y., Wang, C.C., Mineta, K., Ikeo, K., Gojobori, T., Tachibana, M., Kikuchi, T., Kobayashi, T., Shibahara, S., Taya, C., Yonekawa, H., Shiroishi, T., and Yamamoto, H. (2009) Speciﬁc expression of Gsta4 in mouse cochlear melanocytes: a novel role for hearing and melanocyte differentiation. Pigment Cell Melanoma Res., 22, 111–119. Sanchez Hanke, M., Kief, S., Leuwer, R., Koch, U., Moll, I., and Brandner, J.M. (2005) In vitro isolation and cell culture of vestibular inner ear melanocytes. Audiol. Neurootol., 10, 191–200. Mjaatvedt, C.H., Kern, C.B., Norris, R.A., Fairey, S., and Cave, C.L. (2005) Normal distribution of melanocytes in the mouse heart. Anat. Rec. A Discov. Mol. Cell. Evol. Biol., 285, 748–757. Brito, F.C. and Kos, L. (2008) Timeline and distribution of melanocyte precursors in the mouse heart. Pigment Cell Melanoma Res., 21, 464–470. Levin, M.D., Lu, M.M., Petrenko, N.B., Hawkins, B.J., Gupta, T.H., Lang, D., Buckley, P.T., Jochems, J., Liu, F., Spurney, C.F., Yuan, L.J., Jacobson, J.T., Brown, C.B., Huang, L., Beermann, F., Margulies, K.B., Madesh, M., Eberwine, J.H., Epstein, J.A., and Patel, V.V. (2009) Melanocyte-like cells in the heart and pulmonary veins contribute to atrial arrhythmia triggers. J. Clin. Invest., 119, 3420–3436. Yajima, I. and Larue, L. (2008) The location of heart melanocytes is speciﬁed and the level of pigmentation in the heart may correlate with coat color. Pigment Cell Melanoma Res., 21, 471–476. Puig, I., Yajima, I., Bonaventure, J., Delmas, V., and Larue, L. (2009) The tyrosinase promoter is active in a subset of vagal neural crest cells during early development in mice. Pigment Cell Melanoma Res., 22, 331–334. Balani, K., Brito, F.C., Kos, L., and Agarwal, A. (2009) Melanocyte pigmentation stiffens murine cardiac tricuspid valve leaﬂet. J. R. Soc. Interface, 6, 1097–1102.

66 Goldgeier, M.H., Klein, L.E., Klein-

67

68

69

70

71

72

73

Angerer, S., Moellmann, G., and Nordlund, J.J. (1984) The distribution of melanocytes in the leptomeninges of the human brain. J. Invest. Dermatol., 82, 235–238. Fedorow, H., Tribl, F., Halliday, G., Gerlach, M., Riederer, P., and Double, K.L. (2005) Neuromelanin in human dopamine neurons: comparison with peripheral melanins and relevance to Parkinson’s disease. Prog. Neurobiol., 75, 109–124. Zecca, L., Fariello, R., Riederer, P., Sulzer, D., Gatti, A., and Tampellini, D. (2002) The absolute concentration of nigral neuromelanin, assayed by a new sensitive method, increases throughout the life and is dramatically decreased in Parkinson’s disease. FEBS Lett., 510, 216–220. Bush, W.D., Garguilo, J., Zucca, F.A., Albertini, A., Zecca, L., Edwards, G.S., Nemanich, R.J., and Simon, J.D. (2006) The surface oxidation potential of human neuromelanin reveals a spherical architecture with a pheomelanin core and a eumelanin surface. Proc. Natl. Acad. Sci. USA, 103, 14785–14789. Zucca, F.A., Giaveri, G., Gallorini, M., Albertini, A., Toscani, M., Pezzoli, G., Lucius, R., Wilms, H., Sulzer, D., Ito, S., Wakamatsu, K., and Zecca, L. (2004) The neuromelanin of human substantia nigra: physiological and pathogenic aspects. Pigment Cell Res., 17, 610–617. Zecca, L., Tampellini, D., Gatti, A., Crippa, R., Eisner, M., Sulzer, D., Ito, S., Fariello, R., and Gallorini, M. (2002) The neuromelanin of human substantia nigra and its interaction with metals. J. Neural Transm., 109, 663–672. Sulzer, D., Bogulavsky, J., Larsen, K.E., Behr, G., Karatekin, E., Kleinman, M.H., Turro, N., Krantz, D., Edwards, R.H., Greene, L.A., and Zecca, L. (2000) Neuromelanin biosynthesis is driven by excess cytosolic catecholamines not accumulated by synaptic vesicles. Proc. Natl. Acad. Sci. USA, 97, 11869–11874. Zecca, L., Bellei, C., Costi, P., Albertini, A., Monzani, E., Casella, L., Gallorini, M., Bergamaschi, L., Moscatelli, A., Turro, N.J., Eisner, M., Crippa, P.R., Ito,

55

56

2 Classical and Nonclassical Melanocytes in Vertebrates

74

75

76

77

78

79

80

81

82

83

S., Wakamatsu, K., Bush, W.D., Ward, W.C., Simon, J.D., and Zucca, F.A. (2008) New melanic pigments in the human brain that accumulate in aging and block environmental toxic metals. Proc. Natl. Acad. Sci. USA, 105, 17567–17572. Zecca, L., Casella, L., Albertini, A., Bellei, C., Zucca, F.A., Engelen, M., Zadlo, A., Szewczyk, G., Zareba, M., and Sarna, T. (2008) Neuromelanin can protect against iron-mediated oxidative damage in system modeling iron overload of brain aging and Parkinson’s disease. J. Neurochem., 106, 1866–1875. Randhawa, M., Huff, T., Valencia, J.C., Younossi, Z., Chandhoke, V., Hearing, V.J., and Baranova, A. (2009) Evidence for the ectopic synthesis of melanin in human adipose tissue. FASEB J., 23, 835–843. Marusich, M.F. and Weston, J.A. (1991) Development of the neural crest. Curr. Opin. Genet. Dev., 1, 221–229. Le Douarin, N. and Kalcheim, C. (1999) The Neural Crest, 2nd edn, Cambridge University Press, Cambridge. Serbedzija, G.N., Fraser, S.E., and Bronner-Fraser, M. (1990) Pathways of trunk neural crest cell migration in the mouse embryo as revealed by vital dye labelling. Development, 108, 605–612. Mayer, T.C. (1973) Site of gene action in steel mice: analysis of the pigment defect by mesoderm–ectoderm recombinations. J. Exp. Zool., 184, 345–352. Yoshida, H., Kunisada, T., Kusakabe, M., Nishikawa, S., and Nishikawa, S.I. (1996) Distinct stages of melanocyte differentiation revealed by analysis of nonuniform pigmentation patterns. Development, 122, 1207–1214. Jin, E.J., Erickson, C.A., Takada, S., and Burrus, L.W. (2001) Wnt and BMP signaling govern lineage segregation of melanocytes in the avian embryo. Dev. Biol., 233, 22–37. Dorsky, R.I., Moon, R.T., and Raible, D.W. (1998) Control of neural crest cell fate by the Wnt signalling pathway. Nature, 396, 370–373. Larue, L., Kumasaka, M., and Goding, C.R. (2003) Beta-catenin in the

84

85

86

87

88

89

90

91

92

melanocyte lineage. Pigment Cell Res., 16, 312–317. Hornyak, T.J., Hayes, D.J., Chiu, L.Y., and Ziff, E.B. (2001) Transcription factors in melanocyte development: distinct roles for Pax-3 and Mitf. Mech. Dev., 101, 47–59. Watanabe, A., Takeda, K., Ploplis, B., and Tachibana, M. (1998) Epistatic relationship between Waardenburg syndrome genes MITF and PAX3. Nat. Genet., 18, 283–286. Kos, R., Reedy, M.V., Johnson, R.L., and Erickson, C.A. (2001) The winged-helix transcription factor FoxD3 is important for establishing the neural crest lineage and repressing melanogenesis in avian embryos. Development, 128, 1467–1479. Thomas, A.J. and Erickson, C.A. (2009) FOXD3 regulates the lineage switch between neural crest-derived glial cells and pigment cells by repressing MITF through a non-canonical mechanism. Development, 136, 1849–1858. Nishikawa, S., Kusakabe, M., Yoshinaga, K., Ogawa, M., Hayashi, S., Kunisada, T., Era, T., and Sakakura, T. (1991) In utero manipulation of coat color formation by a monoclonal anti-c-kit antibody: two distinct waves of c-kit-dependency during melanocyte development. EMBO J., 10, 2111–2118. Mackenzie, M.A., Jordan, S.A., Budd, P.S., and Jackson, I.J. (1997) Activation of the receptor tyrosine kinase Kit is required for the proliferation of melanoblasts in the mouse embryo. Dev. Biol., 192, 99–107. Cable, J., Jackson, I.J., and Steel, K.P. (1995) Mutations at the W locus affect survival of neural crest-derived melanocytes in the mouse. Mech. Dev., 50, 139–150. Wehrle-Haller, B. and Weston, J.A. (1999) Altered cell-surface targeting of stem cell factor causes loss of melanocyte precursors in Steel17H mutant mice. Dev. Biol., 210, 71–86. Hemesath, T.J., Price, E.R., Takemoto, C., Badalian, T., and Fisher, D.E. (1998) MAP kinase links the transcription factor Microphthalmia to c-Kit signalling in melanocytes. Nature, 391, 298–301.

References 93 Reid, K., Turnley, A.M., Maxwell, G.D.,

94

95

96

97

98

99

100

101

Kurihara, Y., Kurihara, H., Bartlett, P.F., and Murphy, M. (1996) Multiple roles for endothelin in melanocyte development: regulation of progenitor number and stimulation of differentiation. Development, 122, 3911–3919. Lecoin, L., Sakurai, T., Ngo, M.T., Abe, Y., Yanagisawa, M., and Le Douarin, N.M. (1998) Cloning and characterization of a novel endothelin receptor subtype in the avian class. Proc. Natl. Acad. Sci. USA, 95, 3024–3029. Shin, M.K., Levorse, J.M., Ingram, R.S., and Tilghman, S.M. (1999) The temporal requirement for endothelin receptor-B signalling during neural crest development. Nature, 402, 496–501. Hou, L., Pavan, W.J., Shin, M.K., and Arnheiter, H. (2004) Cell-autonomous and cell non-autonomous signaling through endothelin receptor B during melanocyte development. Development, 131, 3239–3247. Nataf, V., Amemiya, A., Yanagisawa, M., and Le Douarin, N.M. (1998) The expression pattern of endothelin 3 in the avian embryo. Mech. Dev., 73, 217–220. Opdecamp, K., Kos, L., Arnheiter, H., and Pavan, W.J. (1998) Endothelin signalling in the development of neural crest-derived melanocytes. Biochem. Cell Biol., 76, 1093–1099. Dupin, E. and Le Douarin, N.M. (2003) Development of melanocyte precursors from the vertebrate neural crest. Oncogene, 22, 3016–3023. Steingrimsson, E., Copeland, N.G., and Jenkins, N.A. (2004) Melanocytes and the microphthalmia transcription factor network. Annu. Rev. Genet., 38, 365–411. McGill, G.G., Horstmann, M., Widlund, H.R., Du, J., Motyckova, G., Nishimura, E.K., Lin, Y.L., Ramaswamy, S., Avery, W., Ding, H.F., Jordan, S.A., Jackson, I.J., Korsmeyer, S.J., Golub, T.R., and Fisher, D.E. (2002) Bcl2 regulation by the melanocyte master regulator Mitf modulates lineage survival and melanoma cell viability. Cell, 109, 707–718.

102 Loercher, A.E., Tank, E.M., Delston,

103

104

105

106

107

108

109

110

R.B., and Harbour, J.W. (2005) MITF links differentiation with cell cycle arrest in melanocytes by transcriptional activation of INK4A. J. Cell Biol., 168, 35–40. Carreira, S., Goodall, J., Aksan, I., La Rocca, S.A., Galibert, M.D., Denat, L., Larue, L., and Goding, C.R. (2005) Mitf cooperates with Rb1 and activates p21Cip1 expression to regulate cell cycle progression. Nature, 433, 764–769. Potterf, S.B., Furumura, M., Dunn, K.J., Arnheiter, H., and Pavan, W.J. (2000) Transcription factor hierarchy in Waardenburg syndrome: regulation of MITF expression by SOX10 and PAX3. Hum. Genet., 107, 1–6. Ludwig, A., Rehberg, S., and Wegner, M. (2004) Melanocyte-speciﬁc expression of dopachrome tautomerase is dependent on synergistic gene activation by the Sox10 and Mitf transcription factors. FEBS Lett., 556, 236–244. Nishimura, E.K., Jordan, S.A., Oshima, H., Yoshida, H., Osawa, M., Moriyama, M., Jackson, I.J., Barrandon, Y., Miyachi, Y., and Nishikawa, S. (2002) Dominant role of the niche in melanocyte stem-cell fate determination. Nature, 416, 854–860. Nishimura, E.K., Granter, S.R., and Fisher, D.E. (2005) Mechanisms of hair graying: incomplete melanocyte stem cell maintenance in the niche. Science, 307, 720–724. Adameyko, I., Lallemend, F., Aquino, J.B., Pereira, J.A., Topilko, P., Muller, T., Fritz, N., Beljajeva, A., Mochii, M., Liste, I., Usoskin, D., Suter, U., Birchmeier, C., and Ernfors, P. (2009) Schwann cell precursors from nerve innervation are a cellular origin of melanocytes in skin. Cell, 139, 366–379. Aoki, H., Yamada, Y., Hara, A., and Kunisada, T. (2009) Two distinct types of mouse melanocyte: differential signaling requirement for the maintenance of non-cutaneous and dermal versus epidermal melanocytes. Development, 136, 2511–2521. Bharti, K., Nguyen, M.T., Skuntz, S., Bertuzzi, S., and Arnheiter, H. (2006) The other pigment cell: speciﬁcation

57

58

2 Classical and Nonclassical Melanocytes in Vertebrates

111

112

113

114

115

116

117

118

119

and development of the pigmented epithelium of the vertebrate eye. Pigment Cell Res., 19, 380–394. Baumer, N., Marquardt, T., Stoykova, A., Spieler, D., Treichel, D., Ashery-Padan, R., and Gruss, P. (2003) Retinal pigmented epithelium determination requires the redundant activities of Pax2 and Pax6. Development, 130, 2903–2915. Fuhrmann, S., Levine, E.M., and Reh, T.A. (2000) Extraocular mesenchyme patterns the optic vesicle during early eye development in the embryonic chick. Development, 127, 4599–4609. Amato, M.A., Arnault, E., and Perron, M. (2004) Retinal stem cells in vertebrates: parallels and divergences. Int. J. Dev. Biol., 48, 993–1001. Perron, M., Boy, S., Amato, M.A., Viczian, A., Koebernick, K., Pieler, T., and Harris, W.A. (2003) A novel function for Hedgehog signalling in retinal pigment epithelium differentiation. Development, 130, 1565–1577. Zhang, X.M. and Yang, X.J. (2001) Temporal and spatial effects of Sonic hedgehog signaling in chick eye morphogenesis. Dev. Biol., 233, 271–290. Nguyen, M. and Arnheiter, H. (2000) Signaling and transcriptional regulation in early mammalian eye development: a link between FGF and MITF. Development, 127, 3581–3591. Hodgkinson, C.A., Moore, K.J., Nakayama, A., Steingrimsson, E., Copeland, N.G., Jenkins, N.A., and Arnheiter, H. (1993) Mutations at the mouse microphthalmia locus are associated with defects in a gene encoding a novel basic-helix-loop-helixzipper protein. Cell, 74, 395–404. Nakayama, A., Nguyen, M.T., Chen, C.C., Opdecamp, K., Hodgkinson, C.A., and Arnheiter, H. (1998) Mutations in microphthalmia, the mouse homolog of the human deafness gene MITF, affect neuroepithelial and neural crest-derived melanocytes differently. Mech. Dev., 70, 155–166. Mui, S.H., Hindges, R., O’Leary, D.D., Lemke, G., and Bertuzzi, S. (2002) The homeodomain protein Vax2 patterns the

120

121

122

123

124

125

126

127

128

129

130

131

132

dorsoventral and nasotemporal axes of the eye. Development, 129, 797–804. Mui, S.H., Kim, J.W., Lemke, G., and Bertuzzi, S. (2005) Vax genes ventralize the embryonic eye. Genes Dev., 19, 1249–1259. Lister, J.A. (2002) Development of pigment cells in the zebraﬁsh embryo. Microsc. Res. Tech., 58, 435–441. Kelsh, R.N., Schmid, B., and Eisen, J.S. (2000) Genetic analysis of melanophore development in zebraﬁsh embryos. Dev. Biol., 225, 277–293. Marks, M.S. and Seabra, M.C. (2001) The melanosome: membrane dynamics in black and white. Nat. Rev. Mol. Cell Biol., 2, 738–748. Jimbow, K. (1999) Biological role of tyrosinase-related protein and its relevance to pigmentary disorders (vitiligo vulgaris). J. Dermatol., 26, 734–737. Orlow, S.J. (1995) Melanosomes are specialized members of the lysosomal lineage of organelles. J. Invest. Dermatol., 105, 3–7. Mottaz, J.H. and Zelickson, A.S. (1967) Melanin transfer: a possible phagocytic process. J. Invest. Dermatol., 49, 605–610. Cohen, J. and Szabo, G. (1968) Study of pigment donation in vitro. Exp. Cell Res., 50, 418–434. Klaus, S.N. (1969) Post-transfer digestion of melanosome complexes and saltatory movement of melanin granules within mammalian epidermal cells. J. Invest. Dermatol., 53, 440–444. Wolff, K., Jimbow, K., and Fitzpatrick, T.B. (1974) Experimental pigment donation in vivo. J. Ultrastruct. Res., 47, 400–419. Hirokawa, N., Noda, Y., and Okada, Y. (1998) Kinesin and dynein superfamily proteins in organelle transport and cell division. Curr. Opin. Cell Biol., 10, 60–73. Mermall, V., Post, P.L., and Mooseker, M.S. (1998) Unconventional myosins in cell movement, membrane trafﬁc, and signal transduction. Science, 279, 527–533. Hara, M., Yaar, M., Byers, H.R., Goukassian, D., Fine, R.E., Gonsalves, J., and Gilchrest, B.A. (2000) Kinesin

References

133

134

135

136

137

138

139

140

141

participates in melanosomal movement along melanocyte dendrites. J. Invest. Dermatol., 114, 438–443. Mehta, A.D., Rock, R.S., Rief, M., Spudich, J.A., Mooseker, M.S., and Cheney, R.E. (1999) Myosin-V is a processive actin-based motor. Nature, 400, 590–593. Wu, X., Bowers, B., Rao, K., Wei, Q., and Hammer, J.A., 3rd (1998) Visualization of melanosome dynamics within wild-type and dilute melanocytes suggests a paradigm for myosin V function in vivo. J. Cell Biol., 143, 1899–1918. Miyamura, Y., Coelho, S.G., Wolber, R., Miller, S.A., Wakamatsu, K., Zmudzka, B.Z., Ito, S., Smuda, C., Passeron, T., Choi, W., Batzer, J., Yamaguchi, Y., Beer, J.Z., and Hearing, V.J. (2007) Regulation of human skin pigmentation and responses to ultraviolet radiation. Pigment Cell Res., 20, 2–13. Thong, H.Y., Jee, S.H., Sun, C.C., and Boissy, R.E. (2003) The patterns of melanosome distribution in keratinocytes of human skin as one determining factor of skin colour. Br. J. Dermatol., 149, 498–505. Hadley, M.E. and Quevedo, W.C., Jr (1966) Vertebrate epidermal melanin unit. Nature, 209, 1334–1335. Hirobe, T. (2005) Role of keratinocytederived factors involved in regulating the proliferation and differentiation of mammalian epidermal melanocytes. Pigment Cell Res., 18, 2–12. Weiner, L., Han, R., Scicchitano, B.M., Li, J., Hasegawa, K., Grossi, M., Lee, D., and Brissette, J.L. (2007) Dedicated epithelial recipient cells determine pigmentation patterns. Cell, 130, 932–942. Marthinuss, J., Andrade-Gordon, P., and Seiberg, M. (1995) A secreted serine protease can induce apoptosis in Pam212 keratinocytes. Cell Growth Differ., 6, 807–816. Santulli, R.J., Derian, C.K., Darrow, A.L., Tomko, K.A., Eckardt, A.J., Seiberg, M., Scarborough, R.M., and AndradeGordon, P. (1995) Evidence for the presence of a protease-activated receptor distinct from the thrombin receptor in

142

143

144

145

146

147

148

149

150

human keratinocytes. Proc. Natl. Acad. Sci. USA, 92, 9151–9155. Nystedt, S., Emilsson, K., Larsson, A.K., Strombeck, B., and Sundelin, J. (1995) Molecular cloning and functional expression of the gene encoding the human proteinase-activated receptor 2. Eur. J. Biochem., 232, 84–89. Nystedt, S., Larsson, A.K., Aberg, H., and Sundelin, J. (1995) The mouse proteinase-activated receptor-2 cDNA and gene. Molecular cloning and functional expression. J. Biol. Chem., 270, 5950–5955. Bohm, S.K., Khitin, L.M., Grady, E.F., Aponte, G., Payan, D.G., and Bunnett, N.W. (1996) Mechanisms of desensitization and resensitization of proteinase-activated receptor-2. J. Biol. Chem., 271, 22003–22016. Sharlow, E.R., Paine, C.S., Babiarz, L., Eisinger, M., Shapiro, S., and Seiberg, M. (2000) The protease-activated receptor-2 upregulates keratinocyte phagocytosis. J. Cell Sci., 113, 3093–3101. Seiberg, M., Paine, C., Sharlow, E., Andrade-Gordon, P., Costanzo, M., Eisinger, M., and Shapiro, S.S. (2000) Inhibition of melanosome transfer results in skin lightening. J. Invest. Dermatol., 115, 162–167. Seiberg, M., Paine, C., Sharlow, E., Andrade-Gordon, P., Costanzo, M., Eisinger, M., and Shapiro, S.S. (2000) The protease-activated receptor 2 regulates pigmentation via keratinocyte– melanocyte interactions. Exp. Cell Res., 254, 25–32. Scott, G. and Zhao, Q. (2001) Rab3a and SNARE proteins: potential regulators of melanosome movement. J. Invest. Dermatol., 116, 296–304. Scott, G., Leopardi, S., Parker, L., Babiarz, L., Seiberg, M., and Han, R. (2003) The proteinase-activated receptor-2 mediates phagocytosis in a Rho-dependent manner in human keratinocytes. J. Invest. Dermatol., 121, 529–541. Scott, G., Leopardi, S., Printup, S., Malhi, N., Seiberg, M., and Lapoint, R. (2004) Proteinase-activated receptor-2 stimulates prostaglandin production in

59

60

2 Classical and Nonclassical Melanocytes in Vertebrates

151

152

153

154

155

156

157

158

159

keratinocytes: analysis of prostaglandin receptors on human melanocytes and effects of PGE2 and PGF2alpha on melanocyte dendricity. J. Invest. Dermatol., 122, 1214–1224. Hanson, D. and DeLeo, V. (1990) Long-wave ultraviolet light induces phospholipase activation in cultured human epidermal keratinocytes. J. Invest. Dermatol., 95, 158–163. Pentland, A.P., Mahoney, M., Jacobs, S.C., and Holtzman, M.J. (1990) Enhanced prostaglandin synthesis after ultraviolet injury is mediated by endogenous histamine stimulation. A mechanism for irradiation erythema. J. Clin. Invest., 86, 566–574. Cardinali, G., Ceccarelli, S., Kovacs, D., Aspite, N., Lotti, L.V., Torrisi, M.R., and Picardo, M. (2005) Keratinocyte growth factor promotes melanosome transfer to keratinocytes. J. Invest. Dermatol., 125, 1190–1199. Marchese, C., Maresca, V., Cardinali, G., Belleudi, F., Ceccarelli, S., Bellocci, M., Frati, L., Torrisi, M.R., and Picardo, M. (2003) UVB-induced activation and internalization of keratinocyte growth factor receptor. Oncogene, 22, 2422–2431. Belleudi, F., Leone, L., Aimati, L., Stirparo, M.G., Cardinali, G., Marchese, C., Frati, L., Picardo, M., and Torrisi, M.R. (2006) Endocytic pathways and biological effects induced by UVBdependent or ligand-dependent activation of the keratinocyte growth factor receptor. FASEB J., 20, 395–397. Boissy, R.E. (2003) Melanosome transfer to and translocation in the keratinocyte. Exp. Dermatol., 12 (Suppl. 2), 5–12. Cardinali, G., Bolasco, G., Aspite, N., Lucania, G., Lotti, L.V., Torrisi, M.R., and Picardo, M. (2008) Melanosome transfer promoted by keratinocyte growth factor in light and dark skin-derived keratinocytes. J. Invest. Dermatol., 128, 558–567. Takeichi, M. (1991) Cadherin cell adhesion receptors as a morphogenetic regulator. Science, 251, 1451–1455. Hirai, Y., Nose, A., Kobayashi, S., and Takeichi, M. (1989) Expression and role of E- and P-cadherin adhesion molecules

160

161

162

163

164

165

166

167

in embryonic histogenesis. II. Skin morphogenesis. Development, 105, 271–277. Tang, A., Eller, M.S., Hara, M., Yaar, M., Hirohashi, S., and Gilchrest, B.A. (1994) E-cadherin is the major mediator of human melanocyte adhesion to keratinocytes in vitro. J. Cell Sci., 107, 983–992. Hakuno, M., Shimizu, H., Akiyama, M., Amagai, M., Wahl, J.K., Wheelock, M.J., and Nishikawa, T. (2000) Dissociation of intra- and extracellular domains of desmosomal cadherins and E-cadherin in Hailey–Hailey disease and Darier’s disease. Br. J. Dermatol., 142, 702–711. Minwalla, L., Zhao, Y., Cornelius, J., Babcock, G.F., Wickett, R.R., Le Poole, I.C., and Boissy, R.E. (2001) Inhibition of melanosome transfer from melanocytes to keratinocytes by lectins and neoglycoproteins in an in vitro model system. Pigment Cell Res., 14, 185–194. Condaminet, B., Redziniak, G., Monsigny, M., and Kieda, C. (1997) Ultraviolet rays induced expression of lectins on the surface of a squamous carcinoma keratinocyte cell line. Exp. Cell Res., 232, 216–224. Jimbow, K., Pathak, M.A., and Fitzpatrick, T.B. (1973) Effect of ultraviolet on the distribution pattern of microﬁlaments and microtubules and on the nucleus in human melanocytes. Yale J. Biol. Med., 46, 411–426. Greatens, A., Hakozaki, T., Koshoffer, A., Epstein, H., Schwemberger, S., Babcock, G., Bissett, D., Takiwaki, H., Arase, S., Wickett, R.R., and Boissy, R.E. (2005) Effective inhibition of melanosome transfer to keratinocytes by lectins and niacinamide is reversible. Exp. Dermatol., 14, 498–508. Jahn, R. and Sudhof, T.C. (1999) Membrane fusion and exocytosis. Annu. Rev. Biochem., 68, 863–911. Osen-Sand, A., Catsicas, M., Staple, J.K., Jones, K.A., Ayala, G., Knowles, J., Grenningloh, G., and Catsicas, S. (1993) Inhibition of axonal growth by SNAP-25 antisense oligonucleotides in vitro and in vivo. Nature, 364, 445–448.

References 168 Araki, S., Tamori, Y., Kawanishi, M.,

Shinoda, H., Masugi, J., Mori, H., Niki, T., Okazawa, H., Kubota, T., and Kasuga, M. (1997) Inhibition of the binding of SNAP-23 to syntaxin 4 by Munc18c. Biochem. Biophys. Res. Commun., 234, 257–262. 169 Virador, V.M., Muller, J., Wu, X., Abdel-Malek, Z.A., Yu, Z.X., Ferrans, V.J., Kobayashi, N., Wakamatsu, K., Ito, S., Hammer, J.A., and Hearing, V.J. (2002) Inﬂuence of alpha-melanocytestimulating hormone and ultraviolet radiation on the transfer of melanosomes to keratinocytes. FASEB J., 16, 105–107. 170 Johannes, L., Lledo, P.M., Roa, M., Vincent, J.D., Henry, J.P., and Darchen, F. (1994) The GTPase Rab3a negatively controls calcium-dependent exocytosis in neuroendocrine cells. EMBO J., 13, 2029–2037. 171 Annaert, W.G., Partoens, P., Slembrouck, D., Bakker, A., Jacob, W., and De Potter, W.P. (1997) Rab3 dissociation and clathrin-mediated endocytosis, two key steps in the exo-endocytotic pathway of large dense-cored vesicles in primary cultures

172

173

174

175

176

of superior cervical ganglia. Eur. J. Cell Biol., 74, 217–229. Geppert, M., Goda, Y., Stevens, C.F., and Sudhof, T.C. (1997) The small GTP-binding protein Rab3A regulates a late step in synaptic vesicle fusion. Nature, 387, 810–814. Araki, K., Horikawa, T., Chakraborty, A.K., Nakagawa, K., Itoh, H., Oka, M., Funasaka, Y., Pawelek, J., and Ichihashi, M. (2000) Small Gtpase rab3A is associated with melanosomes in melanoma cells. Pigment Cell Res., 13, 332–336. Yamamoto, O. and Bhawan, J. (1994) Three modes of melanosome transfers in Caucasian facial skin: hypothesis based on an ultrastructural study. Pigment Cell Res., 7, 158–169. Minwalla, L., Zhao, Y., Le Poole, I.C., Wickett, R.R., and Boissy, R.E. (2001) Keratinocytes play a role in regulating distribution patterns of recipient melanosomes in vitro. J. Invest. Dermatol., 117, 341–347. Lin, J.Y. and Fisher, D.E. (2007) Melanocyte biology and skin pigmentation. Nature, 445, 843–850.

61

wwwwwww

63

3 Biological Chemistry of o-Quinones Patrick A. Riley, Christopher A. Ramsden, and Edward J. Land

3.1 General Biological Signiﬁcance of o-Quinones

An evolutionary approach to the topic of melanins prompts some interesting considerations, for it is clear that, if we adopt an inclusive deﬁnition of melanins (i.e., pigments derived from quinone precursors), these compounds arose early in evolution, and are very widespread both in the animal and plant kingdoms. On the other hand, the specialized organelles in which melanin is synthesized and distributed in vertebrates (the melanosomes) are of much more recent evolutionary origin. While this chronological discrepancy poses many biologically signiﬁcant questions we take the view that the purposes fulﬁlled by melanosomes remain closely bound to the physicochemical properties resulting from the structure of quinone-derived pigments. Quinones are of central importance in biochemical systems, particularly because of their ability to undergo facile one-electron redox reactions, and p-quinones, by virtue of their relative stability, feature among the electron transporters that permit electron movements across lipid membranes (e.g., ubiquinone). By contrast, oquinones are relatively less stable on account of their susceptibility to nucleophilic addition reactions and this property makes them valuable as cross-linking agents. As a result of their reactivity o-quinones possess many biologically signiﬁcant functions, as outlined in the following sections. 3.1.1 Antibiosis

The ability of o-quinones to undergo facile reactions with nucleophiles such as thiol and amino groups (see Section 3.2.2) has been utilized in plants both as a means of antibiosis by damaging invasive organisms (“cytotoxic” action), and as a way of modifying and hardening the protective exterior layers such as are found in seed envelopes (“cross-linking” action). The participation of quinone formation in the “stress responses” of plants is exempliﬁed by the browning reactions in

Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

64

3 Biological Chemistry of o-Quinones

potatoes and bananas that takes place following any surface damage. The browning of fruits appears to be a reaction to invasion by parasitic organisms as well as a generalized response to surface trauma permitting the access of melanogenic substrates (such as dopamine) to catechol oxidases that are normally separately stored in segregated intracellular compartments. As well as exerting an antibiotic action by attacking surface structures the localized production of quinones may have an action in cross-linking structural molecules and rendering them resistant to degradation. This structural reinforcement appears to be the major function of quinonoid polymers in the cell walls of spores. 3.1.2 Defensive Secretions

In insects the cytotoxic potential of o-quinone generation is the basis of their immune system and quinones form an important component of defensive secretions. Quinones have been shown to be a signiﬁcant component of defensive secretions of many small insects such as millipedes and Bombardier beetles. The red-legged millipede (Metiche tanganyiense) is able to prevent predation by the dwarf mongoose with a hot quinone-containing spray. The irritant properties of quinones have been recognized for millennia and they have been employed as laxatives for over 4000 years, and concoctions of madder as a protection against witchcraft have a history extending back to the time of the Essenes [1]. It is likely that the ink of cephalopods exerts its action not so much by obscuring vision (which is of limited signiﬁcance in guiding predators in deep waters) as by the deterrent action on sensitive chemoreceptor organelles of reactive o-quinones generated in the ink, which contains a mixture of tyrosinase and substrate molecules. 3.1.3 Balanid Adhesion

An interesting example of the secretion of tyrosinase utilizing the action of the enzyme on extracellular substrates, in particular on proteins or peptides containing tyrosine residues, is the involvement of quinone cross-linking in adhesion of barnacles. The cements involved in adhesion of several species of barnacle include tyrosine-rich peptides: the protein glues of the blue mussel have been characterized and are members of a 3,4-dihydroxyphenylalanine (dopa)-rich family of polypeptides that cross-link by an oxidative tanning process. Mussels produce protein “byssus” threads secreted by the mussel foot that anchors the mussel to solid substrata in the inter-tidal region. These threads consist of collagen ﬁbers embedded in a polyphenolic protein matrix in which there is a high level of posttranslational hydroxylation of tyrosine and proline residues to give dopa and hydroxyproline, respectively. On oxidation to the corresponding o-quinone of the dopa residues, these proteins form cross-links that stabilize the adhesion. Similar substrate peptides have been isolated in several species of balanids and the crosslinking reactions are tyrosinase-catalyzed [2].

3.1 General Biological Signiﬁcance of o-Quinones

3.1.4 Cuticular Hardening in Insects

The cross-linking action of o-quinones is involved in insect cuticular hardening. The process has been studied quite extensively and relies on the ability of certain side-chain substituted substrate molecules to be successively oxidized so as to form cross-links between adjacent proteins [3]. 3.1.5 Pigmentation

The previous examples are what may be termed initial beneﬁts of melanogenesis in that they derive from the primary oxidation that leads to the production of pigment. The phases of the evolution of melanin pigmentation have been summarized [4] as involving the following steps: i)

Secretion of an o-quinone-generating enzyme with defensive and crosslinking potential.

ii) Intracellular retention of the enzyme with accumulation of a polymerized product of o-quinones and their derivatives, permitting photoreceptor screening. iii)

Limitation of pigment production to speciﬁc cells leading to organismal patterning with roles in camouﬂage and display.

iv)

Transference of pigment to acceptor cells enabling regional effects on durability, photoprotection, and excretion.

Some of these actions are dependent on the structure and properties of melanin. For example, the increased durability of pigmented tissues, demonstrated by the greater resistance to mechanical damage and greater strength of black compared to white feathers, derives from the polymeric structure of melanin [5]. The detailed structure of melanin is unknown, but there is overwhelming evidence that the basic structure entails a backbone of indole moieties that contains both quinone and hydroquinone residues [6]. A high degree of conjugation exists in the polymer which imbues it with strong photon absorption in the visible and UV spectrum. An interesting feature of melanins is the modiﬁcation of optical absorbance characteristics on exposure to light due to oxidation of catecholic groups to quinones. This has the effect of increasing the degree of conjugation of the polymer and the electron delocalization reduces the energy gap between bonding and antibonding π-orbitals. The effect of this is that, as the degree of conjugation increases, lower and lower quantal energies are required for absorption. This bathochromic characteristic of melanin results in signiﬁcant photon absorption even in the IR portion of the spectrum, and this may permit it to act as a means of thermal absorption in poikilotherms and animals in cold climates where solar radiation constitutes a signiﬁcant factor in energy conservation.

65

3 Biological Chemistry of o-Quinones

66

Photoprotection is widely considered to be the most signiﬁcant action of melanin in humans and there is good evidence that increased epidermal melanization is associated with a signiﬁcantly decreased risk of skin cancer related to sun exposure. The mechanism may involve the absorption of energetic photons that might damage important cellular structures such as DNA and, therefore, constitutes an aspect of screening. Another mechanism of protection by melanin may involve scavenging of free radicals. However, an alternative aspect of the protective action of melanin is to consider its functions to include the elimination of cells that may have undergone deleterious mutation. Thus, a genoprotective mechanism exhibited by melanin may involve the generation of free radicals that are toxic to cells that have been exposed to potentially genotoxic doses of light [7]. Owing to the conjugated structure of melanin there is an equilibrium between the quinone and hydroquinone moieties in the polymer that renders the pigment a facile electron exchanger. It can, thus, act as a free radical generator or scavenger [8]. In addition, the relatively high density of negative surface charge on melanin renders it a cation trap and it has been suggested that this constitutes one of the biological beneﬁts conferred by surface pigment since by desquamation in keratocytes melanin provides an excretory pathway for metals and other cationic materials [9]. Finally, as the most prominent surface pigment of vertebrates and some other classes of animals and plants, melanins are important components of external display and camouﬂage.

3.2 o-Quinone Reactivity 3.2.1 Structure and Reactivity

Although o-quinones (1,2-benzoquinones) can often be isolated as crystalline compounds, they are inherently unstable. This is primarily because the six-membered ring is not aromatic (i.e., it does not contain six π electrons in cyclic conjugation in accord with Hückel’s 4n + 2 rule). The driving force for many of the reactions of o-quinones is the formation of an “aromatic sextet,” which is associated with thermodynamic stability. This is illustrated by the facile reduction of o-quinones 1 by ascorbic acid 2 (and many other reducing agents) to give aromatic catechols 3 (Equation 3.1).

R

O

HO +

O

O HO

1

H CH OH 2

H

1

OH

H HO

R

2

O

+ HO

O

O 3

H

O

1

CH2OH OH

O

(3.1)

3.2 o-Quinone Reactivity

Owing to their instability, little structural information is available for simple monocyclic o-quinones. Stability can be increased by substituent effects and the molecular structure of the sterically hindered 3,5-di-tert-butyl derivative 4 has been determined using X-ray diffraction [10]. In contrast to catechol 5, in which all the C–C bond lengths (average 1.385 Å) are characteristic of an aromatic ring (benzene 1.39 Å)[11], the C–C bonds of the o-quinone 4 have lengths corresponding to either double bonds (average 1.342 Å) or single bonds (range 1.439 to 1.554 Å). Magnetic criteria such as nucleus-independent chemical shifts (o-quinone NICS(1) 1.3; benzene NICS(1) −12.8) and proton chemical shifts also indicate that o-quinones are non-aromatic [12].

O

But 1.340

1.399

HO

1.378 O

HO

But 1.344 4

1.391 5

In addition to reduction to a catechol (e.g., Equation 3.1), there are other mechanisms by which o-quinones react to achieve aromaticity. A common mode of reaction is addition of a nucleophile (Equation 3.2), which also leads to aromatic catechol derivatives (e.g., 6).

O

R1

H2N–R2

O

R1 NH2 R2

–

O

HO

R1

HO

N R2 H

+

O H

6

(3.2)

It is beyond the scope of this chapter to survey all the modes of reactions of oquinones. Only reactions of known biological signiﬁcance are covered in the following sections. However, cycloaddition is a common mode of reaction and the following examples are worthy of brief mention. [4 + 1] Cycloaddition readily occurs with phosphorus derivatives to give the cycloadducts 7 (Equation 3.3), which also achieve aromatic stability. The stable adducts 7 can be useful for characterizing o-quinones [13]. O O

R1

PR3

R1

O R3P

(3.3)

O 7

Another common type of cycloaddition is the [4 + 2] Diels–Alder reaction with alkenes that can form the cycloadducts 8 and/or the cycloadducts 9 (Equation 3.4).

67

68

3 Biological Chemistry of o-Quinones

In some cases an o-quinone will react with itself to give a dimeric cycloadduct. Other types of cycloaddition reaction of o-quinones have been reviewed [13, 14]. R2

R1

O

R2

R1

O

R2 and/or

+ O

R2

R2

R2

O 8

CO CO

R1

9 (3.4)

It should also be noted that because of their inherent reactivity, some o-quinones rapidly react via variations of the above modes of reaction to give unexpected products. Thus, the o-quinone 10, formed from pyrogallol, rapidly dimerizes with elimination of carbon monoxide to give purpurogallin 11 (Equation 3.5) [15]. OH O

OH O

–CO

2

OH

HO

(3.5)

HO

O 10

11

3.2.2 Reduction

Reduction is the process of adding electrons. o-Quinones readily accept one electron to form a semiquinone radical anion 12 followed by a second electron to form a catechol dianion 15 (Scheme 3.1). Depending upon the conditions these species R1

O

e–

• –

O

R1

O

O

1

H+

13 e–

R1

HO –

Scheme 3.1

H+

–

14

R1

O

–

O

3

R1

O

12

R1

HO

•

HO

H•

HO

H+

O 15

3.2 o-Quinone Reactivity Table 3.1

Redox potentials (Eo) of selected quinones.

Entry

Quinone

Eo (V)

Reference

1 2 3 4 5 6 7 8

1,2-benzoquinone 1,4-benzoquinone 2,3,4,5-tetrachloro-1,2-benzoquinone 3,5-di-tert-butyl-1,2-benzoquinone 4-hydroxy-1,2-benzoquinone 1,2-naphthoquinone 1,2-anthraquinone 9,10-phenanthraquinone

0.795 0.711 0.830 0.580 0.594 0.576 0.490 0.460

[16] [17] [16] [18] [19] [20] [20] [20]

may be protonated (13 and 14). In some cases the second reductive step may occur via hydrogen atom transfer from the partially oxidized reducing agent (XHᠨ). This → 3) accounts for many of the biologically signiﬁreversible reductive process (1 ← cant reactions of o-quinones. The ease of reduction of o-quinones to catechols (1 → 3) is indicated by the redox potential (Eo), which can be measured potentiometrically. Substituents, including fused rings, can signiﬁcantly modify the redox potential. As the redox potential increases the quinone becomes a more powerful oxidizing agent. Selected redox potentials are shown in Table 3.1. Electron-withdrawing groups increase the redox potential (e.g., Entry 3) and hence the oxidizing power of the o-quinones, that is, they are more easily reduced. Conversely, electron-donating substituents (e.g., Entries 4 and 5) and fused rings (e.g., Entries 6–8) reduce the oxidizing power. A difference in redox potentials of o-quinones leads to a process known as redox exchange, which plays a signiﬁcant role in the early stages of melanin biosynthesis. This process is summarized in Equation 3.6 in which an o-quinone 17 oxidizes a catechol 16 to give a new o-quinone 18 and is itself reduced to the catechol 19. Thus, an o-quinone with a high redox potential such as 2,3,4,5-tetrachloro-1,2benzoquinone (Entry 3, Table 3.1) will oxidize a wide variety of catechols to the corresponding o-quinones. This process occurs via intermediate semiquinone radicals, as shown in Scheme 3.1, with the catechol acting as the reducing agent. In fact, if semiquinone radicals (e.g., 12 → ← 13) are generated in solution by, for example, pulse radiolysis, they very rapidly disproportionate to a mixture of the corresponding catechol and o-quinone.

R1

OH

R2

OH 16

+

O

R3

O

R4 17

R1

O O

R2 18

+

HO

R3

HO

R4 19 (3.6)

69

70

3 Biological Chemistry of o-Quinones

Scheme 3.2 summarizes the role of redox exchange in the early stages of melanin biosynthesis [21, 22]. Since dopachrome 23 is a quinone with a strongly electron-donating substituent (NH), it has a smaller redox potential than dopaquinone 21. This, therefore, favors redox exchange between cyclodopa 20 and dopaquinone 21 to give dopa 24 by a nonenzymatic mechanism. In a similar way, cysteinyldopa 22 is oxidized to cysteinyldopaquinone 25. For further discussion of the balance between these alternative redox exchange pathways, and their inﬂuence on tyrosinase kinetics, see Section 3.2.3.

HO

CO2–

O

–

CO2 HO

NH2

O

N H

NH2

HO Cys

20 cyclodopa

21 dopaquinone

22 cysteinyldopa redox exchange

redox exchange

O CO2 O

CO2–

HO

Cys

CO2–

HO

–

NH2

HO

N H

CO2–

O

NH2

O Cys

23 dopachrome

24 dopa

25 cysteinyldopaquinone

eumelanin pathway

pheomelanin pathway

Scheme 3.2

o-Quinones with a tertiary amine side-chain 26 cyclize to the betaines 27 that in aqueous solution are in equilibrium with the catechols 28 [23]. As the species 27 and 28 are associated with a strongly electron-withdrawing substituent (R2N+), they would be oxidized to a quinone derivative 29 with a much higher redox potential than the quinone precursor 26. Consequently, in contrast to the cyclodopa– dopaquinone system (Scheme 3.2), redox exchange does not take place between 26 and 27/28 (Scheme 3.3) and this has been used to study the mechanism of activation of tyrosinase in vitro (see Section 3.3) [24, 25]. –

O NR2

O

O

H+ +

HO

N R2 27

26 Scheme 3.3

O

HO +

HO

N R2 28

X

+

N R2

O

29

3.2 o-Quinone Reactivity

3.2.3 Addition Reactions: Intermolecular addition

o-Quinones are electron-deﬁcient molecules and react with nucleophiles such as amines to give addition products (e.g., 6 in Equation 3.2). The example shown in Equation 3.2 is formally a 1,4-addition to a conjugated carbonyl, commonly referred to as Michael addition. Since the nucleophile can attack more than one position of the ring, including the carbonyl carbons, nucleophilic attack can lead to a variety of addition reactions that are largely beyond the scope of this chapter. One important case of intermolecular addition that merits mention is the addition of cysteine to dopaquinone 21 to give cysteinyldopa 22 (Scheme 3.2). This is formally a 1,6-addition to a conjugated carbonyl. Although 5-S-cysteinyldopa 30 is the major product (74%), there is a smaller amount of the alternative 1,6-adduct (2-S-cysteinyldopa 31) (14%) and a tiny amount (1%) of the 1,4-adduct (6-S-cysteinyldopa 32) (Scheme 3.4) [21, 26, 27]. The small amount of the 1,4-adduct 32 is surprising since Michael addition is favorable for amine nucleophiles. The product distribution for cysteine addition has not been rationalized. Amines are hard nucleophiles (charge controlled), whereas thiols are soft nucleophiles (orbital controlled) and it is possible that the nature of the o-quinone frontier molecular orbitals controls the regioselectivity. Alternatively, thiols are reducing agents and addition may take place via a radical mechanism, possibly involving a semiquinone radical.

CO2–

O

NH2

O

21 dopaquinone Cys

CO2–

HO

NH2

HO

+

Cys HO HO

CO2– NH2

+

CO2–

HO HO

Cys

NH2

Cys 30 5-cysteinyldopa 74%

31 2-cysteinyldopa 14%

32 6-cysteinyldopa 1%

Scheme 3.4

3.2.4 Polymerization

A special case of intermolecular addition is the formation of melanin polymer from indole building blocks. Structural studies suggest that each step in the process is an addition to an indolequinone intermediate. In the eumelanin pathway decarboxylation of dopachrome 23 and subsequent tautomerization gives

71

72

3 Biological Chemistry of o-Quinones

5,6-dihydroxyindole (DHI) 33 that is partially oxidized to highly reactive 5,6-indolequinone (IQ) 34 [22, 28]. Product analysis of oligomers suggests that polymerization occurs by nucleophilic attack of DHI 33 on the IQ 34 to give a mixture of intermolecular addition products. The mode of addition seems to favor 2,2′-, 2,4′-, and 2,7′-linkages (i.e., 35–37). This is surprising since indoles usually react with electrophiles at the 3-position. The dihydroxyindole dimers 35–37 then react further in a similar manner giving polymeric material. The dimeric structures in Scheme 3.5 are only representative since DHI-2-carboxylic acid (DHICA) is also known to form polymeric material. 4

HO

dopachrome 23

3

O

2 HO

7

N H

O

N H

33

34

NH HO

2 4′

HO

N H HO

HN HO + HO

OH

35

HO

2 7′

2 2′

+ HO

N H HO

H N

OH OH

N H

OH

36

37

Scheme 3.5

3.2.5 Intramolecular Addition (Cyclization)

In contrast to intermolecular addition, a simpler situation arises when a nucleophile is attached to a side-chain on an o-quinone ring, as is the case for dopaquinone 21. In this case the side-chain limits the rapid nucleophilic attack by the amine to either position 3 or position 5. In practice attack at position 3 is kinetically unfavorable [29] and only the product 20 is formed (Scheme 3.6). The alternative intramolecular cyclization product 38 is not detected. CO2– HO HO

CO2– 5 N H 20 cyclodopa

Scheme 3.6

O O

3

4 5

CO2– NH2

21 dopaquinone

HO X

HN 3

HO

38

3.2 o-Quinone Reactivity

73

It is noteworthy that when the side-chain is lengthened by one carbon atom (i.e., propylamines 40) nucleophilic attack at position 4 to give a spiro product 39 is the fastest reaction (Scheme 3.7). However, this spiro-cyclization is reversible, since the kinetic product 39 is unstable (not aromatic) and the system rapidly equilibrates giving the thermodynamic product 41 [23, 30, 31]. Dopaquinone 21, and related ethylamines, do not undergo spiro-cyclization because four-membered ring formation is unfavorable. +

–

O

3

R2N 4

O

4 5

O

O

–

39

O

HO

NR2

40

+

5 N R2 41

Scheme 3.7

3.2.6 Addition–Elimination (Substitution) Reactions

Addition of a nucleophile to an o-quinone is sometimes followed by elimination of a small molecule, which is often water. The overall reaction is commonly referred to as substitution. An example is the spontaneous cyclization of 5-Scysteinyldopaquinone 42 to the quinonimine 43 with elimination of water (Scheme 3.8). Subsequent tautomerization gives the 1,4-benzothiazine ring 44 characteristic of pheomelanin pigment [21].

CO2–

O

NH2

O H2N

S

CO2–

O

NH2

N H2O

–

O2C

–

O2C

42

CO2–

HO

NH2

N

S

–

O2C

43

S 44

Scheme 3.8

Another example is the cyclization of 6-ﬂuorodopaquinone 45 with elimination of hydrogen ﬂuoride (Equation 3.7), which gives dopachrome 23 directly without requiring oxidation (cf. Scheme 3.2) [32]. CO2–

O O

O

F NH2

O HF

45

CO2– N H

23 dopachrome

(3.7)

74

3 Biological Chemistry of o-Quinones

3.3 Role of o-Quinones in Melanogenesis 3.3.1 Nonenzymatic Formation of Melanogenic Intermediates 3.3.1.1 Contributions from Pulse Radiolysis to the Chemistry of Eumelanogenesis and Pheomelanogenesis Up to the mid-1980s, much of what was known about the initial chemical steps of melanogenesis had been discovered by enzymic oxidation studies in which slow enzyme-catalyzed steps are rate determining, precluding the build-up of high enough concentrations of the reactive intermediates to allow their kinetics to be investigated. This pathway, which shows the principal steps of melanin formation, is summarized in Scheme 3.9, essentially an updated version of the classic Raper– Mason scheme for melanogenesis [33–35]. The conversion of tyrosine 46 to melanin was proposed to involve a series of oxidation steps, the initial being the tyrosinase-catalyzed oxidation of tyrosine to the highly reactive o-quinone dopaquinone, the subsequent reactions leading eventually to the eumelanic and pheomelanic polymers proceeding spontaneously. From the mid-1980s, the application of the technique of pulse radiolysis to this ﬁeld enabled the reaction rate constants associated with a number of the steps shown in Scheme 3.9 to be measured, at the same time providing very strong direct evidence of the accuracy of the commonly accepted scheme of melanin formation. This technique is, for example, able to generate “instantaneously” (i.e., within a few milliseconds) relatively high concentrations (e.g., up to around 10−4 M) of dopaquinone 21. This can be achieved in simple aqueous solution by the careful choice of various solutes including, for example, dopa, KBr, and phosphate buffer, at speciﬁc relative concentrations, all under an atmosphere of nitrous oxide. The technique of pulse radiolysis consists of delivering short pulses of ionizing radiation, typically of the order of up to a few microseconds in duration, from an electron linear accelerator to samples contained in high-purity quartz optical cells. Light, often from a xenon arc lamp, is passed continuously through the sample cell, usually at right angles to the path of the electron beam. The short pulse of radiation results in the formation within the sample of transient species that have characteristic absorption spectra. The remainder of the pulse radiolysis equipment is, in essence, a very fast response spectrophotometer, able to follow changing absorption spectra over timescales ranging from microseconds, or less, to minutes. Most of the relevant melanin precursor studies employed the pulse radiolysis apparatus developed from the 1960s by Keene [36] at the Paterson Laboratories, now the Paterson Institute for Cancer Research, Christie Hospital and Holt Radium Institute, Manchester, UK. In 2000, the detection equipment was transferred to the Synchrotron Radiation Laboratory, Daresbury, Warrington, UK, where it continued to be used with a different linear accelerator until 2008 (Figure 3.1).

3.3 Role of o-Quinones in Melanogenesis

HO

COOH NH2 46

O

COOH

r1

HO

COOH HO

N 20 H

HO

cysteine

NH2

O

r3

COOH NH2

HO

21

S

r2

H2N

r4

COOH

O

O

COOH HO

NH2

HO

23

N 47

dopa quinone

HO

r8

25

S

r5

COOH

H2N

COOH HO

NH2

O

24 HO

HO

COOH

COOH

N

HO

22

COOH

O

N 33

N r9

S

HOOC

43

r6

dopa O

O

COOH

HO

COOH N

O

N

O

48

NH2

34

NH2

N (HOOC)

S

44 r7

Eumelanins

Pheomelanins

Scheme 3.9 Outline of reactions involved in melanogenesis showing the reactions for which rate constants have been determined (see Table 3.2).

When ionizing radiation is incident upon a dilute aqueous solution, the radiation, unlike light, is almost completely absorbed by the solvent (water) generating within it mainly two types of very reactive free radicals, namely hydrated electrons (e−aq) and HOᠨ radicals. If the solution is saturated with N2O, all the e−aq are converted into further HOᠨ radicals. These hydroxyl radicals are highly oxidizing and have a tendency to add to aromatic solutes, as well as causing one-electron oxidations. In the presence of excess KBr, these HOᠨ radicals, from both sources, generate Br2ᠨ− radicals which then cause exclusive one-electron oxidation, in the

75

76

3 Biological Chemistry of o-Quinones

Figure 3.1 The Manchester Pulse Radiolysis apparatus reinstalled at the Daresbury Synchrotron Radiation Laboratory. The reaction chamber is being set up by Professor E.J. Land, Dr Ruth Edge, and Dr Suppiah Navaratnam.

case of dopa yielding dopasemiquinone radicals. The unstable dopasemiquinones then disproportionate within a few milliseconds into dopaquinone and reformed dopa. From measurements made at 380 nm (the wavelength maximum of dopaquinone 21) in the absence of further additives, the dopaquinone decays over hundreds of milliseconds, with the subsequent formation of dopachrome 23, a precursor of eumelanin, with a wavelength maximum of 480 nm. However, the kinetics of formation at 480 nm does not match exactly the decay at 380 nm, there clearly being an intermediate between dopaquinone and dopachrome. This is interpreted in terms of dopaquinone cyclizing over hundreds of milliseconds (k = 3.8 s−1) [37], forming cyclodopa 20 which rapidly cross-reacts with the remaining dopaquinone 21 to yield reformed dopa 24 and dopachrome 23. In order to measure directly the rate constant of redox exchange between dopaquinone and cyclodopa, the moderately stable cyclodopa was synthesized chemically, and the kinetics of formation of dopachrome upon addition of cyclodopa to pulseradiolytically generated dopaquinone followed. The addition of cyclodopa 20 did indeed result in an increase in the rate of formation of dopachrome 23 absorption at 480 nm, leading to a rate constant for redox exchange between dopaquinone 21 and cyclodopa 20 of 5.3 × 106 M−1 s−1. This rate constant, together with the rate constant for the unimolecular decay of dopaquinone, led to a satisfactory simulation of the optical density versus time curves at 380 and 480 nm obtained from the pulse radiolysis of dopa alone [37].

3.3 Role of o-Quinones in Melanogenesis

In the presence of cysteine, dopaquinone 21 undergoes reductive addition to give cysteinyldopa 22, mainly the 5-S isomer 30, thus initiating the pheomelanic pathway. From studies of the enhanced rate of decay of dopaquinone absorption at 380 nm, a rate constant of 3 × 107 M−1 s−1 was obtained for the reaction of cysteine with dopaquinone [38]. Dopaquinone 21 can also be shown by pulse radiolysis to undergo spontaneous redox exchange with 5-S-cysteinyldopa 30, leading to 5-Scysteinyldopaquinone 25. Although the latter, like dopaquinone, has an absorption maximum at 380 nm, its extinction coefﬁcient is over 3 times greater. This enabled the transition from dopaquinone to cysteinyldopaquinone to be followed easily, and the resultant rate constant of 8.8 × 105 M−1 s−1 was obtained [39]. 5-S-Cysteinyldopaquinone 25 was shown to cyclize (k = 10 s−1) spontaneously to a quinonimine 43, wavelength maximum 540 nm, which, in turn, decayed by tautomerization to a benzothiazine 44 (k = 6.0 s−1). The latter was found to decay subsequently with a rate constant of 0.5 s−1 [40]. Later isolatable intermediates in the series of chemical reactions leading to eumelanin are DHICA (47), a rearrangement product of dopachrome, and its decarboxylation product DHI (33). In an analogous way in which the above redox exchanges between dopaquinone and cyclodopa or 5-S-cysteinyldopa were studied, pulse radiolysis investigations were carried out seeking redox exchange between dopaquinone and DHI or DHICA [41]. Evidence for a redox exchange occurring between dopaquinone 21 and DHI 33 with a rate constant of 1.4 × 106 M−1 s−1 was gained, although the identity of the actual DHI quinoid tautomer 34 awaits application of a more structurally informative detection technique, such as time-resolved resonance Raman spectroscopy. The results obtained with mixtures of dopaquinone 21 and DHICA 47 were interpreted as demonstrating that, in this case, redox exchange is around 10 times slower than for dopaquinone to DHI and also goes less to completion [41]. A summary of the rate constants, obtained by pulse radiolysis for the early chemical reactions of eumelanogenesis and pheomelanogenesis, is provided in Table 3.2 (see also [31]).

Table 3.2

Reaction r1 r2 r3 r4 r5 r6 r7 r8 r9

Rate constants for the reactions shown in Scheme 3.9. First order (s−1)

Second order (M−1 s−1)

3.8 5.3 × 106 3.0 × 107 8.8 × 105 10.0 6.0 0.5 1.6 × 105 1.4 × 106

Reference [37] [37] [38] [39] [40] [40] [40] [41] [41]

77

78

3 Biological Chemistry of o-Quinones

3.3.2 Balance between Eumelanogenesis and Pheomelanogenesis

Both eumelanin and pheomelanin are derived from dopaquinone, and the divergent pathways of biogenesis may be described by the balance between the rate constants of the nonenzymatic reactions involved in the initial steps. Thus, taking dopachrome and cysteinyldopaquinone as representative of the divergent pathways, an index of divergence (D), deﬁned as the ratio of the products of the rate constants of the reactions involved, has been proposed [37]: D=

r3 ⋅ r4 ⋅ [cysteine ] r1 ⋅ r2

where r1, r2, and r3 are the rate constants determined by pulse radiolysis for, respectively, dopaquinone cyclization, redox exchange with cyclodopa, and cysteine addition, and r4 is the rate constant for dopaquinone redox exchange with 5-Scysteinyldopa. This leads to a “cross-over value” (i.e., for D = 1) for switching between predominance of eumelanogenesis to predominant pheomelanogenesis, when the cysteine concentration at the site of melanogenesis is 7.6 × 10−7 M [37]. The factors regulating the intramelanosomal cysteine concentration are discussed elsewhere in this volume (see Chapter 6). 3.3.3 Control of Melanogenesis: Phase I Melanogenesis

The initial reactions in the melanogenic pathway (often referred to as phase I melanogenesis) are of signal importance because of their inﬂuence on the activity of tyrosinase and their part in the control of the nature of the ﬁnal pigmentary product (see Scheme 3.10). An important feature of the spontaneous reactions outlined above is their signiﬁcance in generating the catecholic intermediate, dopa. Dopa is central to the control of melanogenesis because of its two actions in inﬂuencing the activity of tyrosinase both as an activator of mettyrosinase by acting as a reducing substrate and as a suicide inactivator of tyrosinase. 3.3.4 Tyrosinase Activation

Tyrosinase is unusual in that it exhibits two catalytic functions: (i) oxygenase activity (“cresolase” action, shown as “t’ase I” in Scheme 3.10) in which monohydric phenols have oxygen substituted at the 2-position giving an o-quinone [42, 43], and (ii) oxidase activity (“catecholase” action, shown as “t’ase II” in Scheme 3.10) [44] in which a catechol is oxidized to the corresponding o-quinone. One of the characteristics of in vitro oxidation of tyrosine 46 is a “lag period” [45]. Native tyrosinase occurs mainly as met-tyrosinase 49, which cannot oxidize phenols (e.g., tyrosine) and needs to be reduced to deoxy-tyrosinase 50 by a catechol before phenol oxida-

3.3 Role of o-Quinones in Melanogenesis tyrosine

t’ase I cysteine

dopaquinone

cyclodopa

cysteinyldopa

t’ase II

dopachrome

dopa

cysteinyldopaquinone

tyrosinase inactivation Eumelanin

Pheomelanin

pheo/eumelanin balance Scheme 3.10

Phase I reactions and the control of melanogenesis.

tion can begin [46]. In vitro this activating catecholic substrate is generated indirectly by fast redox exchange of dopaquinone formed slowly by the small amount of oxy-tyrosinase 51 present in native tyrosinase. The lag period ends when all the enzyme has been activated by this indirect and relatively slow nonenzymatic formation of dopa (Scheme 3.11) [24, 25]. 3.3.5 Tyrosinase Inactivation

Another unusual feature of tyrosinase is that the oxidase pathway exhibits inactivation kinetics [47–53]. It was shown in 1982 by Dietler and Lerch [54] that this inactivation is linearly correlated with the loss of 50% of the copper atoms from the active site, but the mechanism to account for the loss of copper remained elusive. Dietler and Lerch suggested that it might be the result of oxidative modiﬁcation of the histidine residues that coordinate the copper atoms in the active site of the enzyme [54], but radical scavengers were without effect on the inactivation process [51, 54] and it is probable that the histidine oxidation they observed was secondary to the inactivation process.

79

80

3 Biological Chemistry of o-Quinones

oxygenase pathway

N

– N O 2+ N Cu O N –

51 oxy-tyrosinase

R HO O

R

oxidase pathway

Cu2+ N N

HO HO

R

R

O X

O2

O

O +

+

N N

N

N N Cu1+ N

1+

Cu

50 deoxy-tyrosinase

HO HO

R N N

N

H

O 2+ –

Cu

N N Cu2+ N

49 met-tyrosinase

N = histidine ligand Scheme 3.11

Recently a mechanism was proposed to account for the copper loss during tyrosinase inactivation [55]. Brieﬂy stated, this “Quintox mechanism” postulates that oxy-tyrosinase 51 sometimes binds catechol substrates in the oxygenase mode (i.e., as if they were phenols). Scheme 3.12(a) shows the intermediate 52 formed by binding of a phenol to oxy-tyrosinase followed by elimination of an o-quinone 54 and generation of deoxy-tyrosinase 50. If a catechol were to bind to the active site of the enzyme in a similar way (53, Scheme 3.12b), the extra hydroxy substituent could deprotonate leading a 3-hydroxy-o-quinone 55, reductive elimination of Cu(0), and an irreversibly inactivated form of tyrosinase 56. This processing of catechols as phenols in the oxygenase cycle accounts for the observed suicide inactivation and is consistent with a number of experimental studies [56, 57]. A caveat is that the majority of the recent experimental data bearing on the question of suicide inactivation are derived from work with tyrosinase from mushrooms (Agaricus bisporus) and this limits the extent to which the conclusions drawn from them can be generalized. However, the strongly conserved active center of tyrosinases obtained from different sources [58] makes it probable that very similar, if not identical, mechanisms apply to this class of enzyme in general. While there are no studies unequivocally demonstrating the formation of zerovalent copper during the suicide inactivation of tyrosinase, this mechanism seems to be the most plausible explanation for the loss of enzymatic activity and a variation of the reductive–elimination mechanism has also been proposed [59]. The current state of the evidence has been the topic of a recent review [60] and may be summarized as follows.

3.3 Role of o-Quinones in Melanogenesis a)

b)

H O –

N

N

N Cu2+ – N– O O

Cu2+

N

N N

N

O

O

N N

R

53

O

Cu2+

O

O

52

O

N

– Cu2+ –

–

H

R

H O

N

OH

O R

R

54

N N

+ H2O N

N Cu

55

1+

Cu1+

+ N

N

N

N

50 deoxy-tyrosinase

H2 O N N Cu2+ Cu N N

56 inactivated tyrosinase

Scheme 3.12

A strong prediction arising from the mechanistic hypothesis proposed by the Quintox group [55] is that catechol oxidases (EC 1.10.3.1), enzymes closely related to tyrosinase but which lack oxygenase activity due to subtle differences in the active site [61–63], will fail to exhibit suicide-inactivation kinetics since the copper reduction requires oxygenase-catalyzed generation of a hydroxylated intermediate (Scheme 3.12b). This is not predicted by the alternative mechanism recently advanced by Muñoz-Muñoz et al. [59] since their proposal depends on a protonation of the peroxy oxygen in a bidentate catechol complex. The catechol oxidase extracted from bananas (Musa cavendishii) shows no activity towards monohydric phenols and, in contrast to tyrosinase, it oxidizes a number of catecholic substrates without exhibiting suicide-inactivation kinetics [56]. Further indirect evidence of the involvement of an oxygenase presentation in the inactivation mechanism is the inhibitory effect of addition of a monohydric phenol to the catechol oxidation mixture. The relative inactivation rate (the ratio of the inactivation rate to the initial rate of oxidase activity) of 4-methylcatechol oxidation by tyrosinase is diminished with increasing concentrations of 4-methylphenol [55]. According to the Quintox mechanism the inactivation comes about by a proportion of the substrate binding to the active site in an alternative orientation. The catechol may bind either in the usual bidentate oxidase mode to the active-site copper atoms 57 or like a phenol in the oxygenase mode 58. The orientation of

81

82

3 Biological Chemistry of o-Quinones

these binding modes to oxy-tyrosinase differs in respect of the plane deﬁned by the copper atoms and the coordinated molecular oxygen [64–67]. Both modes (57 and 58) have well-deﬁned and inﬂexible locations of the catechol unit and its associated substituents. It might be anticipated, therefore, that structural constraints associated with different binding modes would be reﬂected by a differential inﬂuence of substituents on the rate of oxidation (k1) and the rate of inactivation (k2). Such differences have been observed by quantitative analysis of a series 4-substituted catechols with ring substituents chosen to give a wide variation and minimal correlation of substituent properties [68].

N N

H O

N

N

– Cu2+ –

2+

Cu

O O H O–

–

H N O N 2+ – Cu – N– O O

N

N H

O

N Cu2+

R

N N

N N

H O

N

– Cu2+ –

N Cu2+

O O H O–

–

F

N N

F

R 57

58

59

There is evidence in the literature of the 5-hydroxylation of dopa that conﬁrms the ability of tyrosinase from different sources to process catechols by the oxygenase pathway [69–72] and a strong prediction of the Quintox mechanism is the generation of a hydroxylation product 55 of the catecholic substrate during the inactivation reaction (Scheme 3.12b). As the inactivation reaction is relatively rare and involves small quantities of enzyme, in the experimental system used the o-hydroxylated quinone product 55 is a minor component of the reaction mixture. Nevertheless, the product 55 has been identiﬁed by high-performance liquid chromatography/mass spectrometry in the case of 4-methylcatechol oxidation by tyrosinase and shown to be correlated with the loss of enzyme activity [56]. Since the Quintox suicide-inactivation mechanism requires o-hydroxylation to take place, the inactivation kinetics of tyrosinase during catechol oxidation should be prevented if an unsubstituted ring carbon adjacent to the catecholic function is not available. Since this is not a requirement of the mechanism suggested by Muñoz-Muñoz et al. [59] this provides a critical experimental test of the mechanisms. As predicted by the Quintox mechanism, 3,6-diﬂuorocatechol 59, which cannot act as an oxygenase substrate, acts as an oxidase substrate for tyrosinase, but does not exhibit suicide-inactivation kinetics [57]. The oxygen utilization during the oxidation of 3,6-diﬂuorocatechol 59 by tyrosinase shows ﬁrst-order kinetics and contrasts with the kinetics of inactivating substrates. All the experimental tests so far carried out have been consistent with the predicted outcome of the Quintox inactivation mechanism of tyrosinase.

References

References 1 Steckoll, S.M., Goffer, Z., Haas, N., and

2

3

4

5

6 7

8

9

10

11 12

13

Nathan, H. (1971) Red-stained bones from Qunran. Nature, 231, 469–470. Yamamoto, H. and Tatehata, H. (1995) Oxidative reaction mechanism by use of tyrosinase towards synthetic mytilid bivalve adhesive protein precursors. J. Mar. Biotechnol., 2, 95–100. Sugumaran, M. (1991) Molecular mechanisms for mammalian melanogenesis – comparison with insect cuticular sclerotization. FEBS Lett., 293, 4–9. Riley, P.A. (1995) The evolution of melanogenesis, in Melanin: Its Role in Human Photoprotection (ed. L. Zeise, M.R. Chedekel, and T.B. Fitzpatrick), Valdemar, Overland Park, KS, pp. 11–22. Robins, A.H. (1991) Biological Perspectives on Human Pigmentation, Cambridge University Press, Cambridge. Nicolaus, R.A. (1968) Melanins, Hermann Press, Paris. Riley, P.A. (1974) Melanin and melanocytes, in The Physiology and Pathophysiology of the Skin, vol. 3 (ed. A. Jarrett), Academic Press, London, pp. 1104–1127. Sichel, G., Corsaro, C., Scalia, M., Sciuto, S., and Geremia, E. (1987) Relationship between melanin content and superoxide dismutase (SOD) activity in the liver of various species of animals. Cell Biochem. Funct., 5, 123–128. Riley, P.A. (1997) Epidermal melanin: sun screen or waste disposal? Biologist, 44, 408–411. Horng, D.-N., Chyn, J.-P., Shieh, K.-J., Chou, J.-L., and Wen, Y.-S. (1999) 3,5-Di-tert-butyl-1,2-benzoquinone. Acta Crystallogr. C, 55, 652–653. Brown, C.J. (1966) The crystal structure of catechol. Acta Crystallogr., 21, 170–174. Golas, E., Lewars, E., and Liebman, J.F. (2009) The quinones of benzocyclobutadiene: a computational study. J. Phys. Chem. A, 113, 9485–9500. Ramsden, C.A. (2010) Heterocycleforming reactions of 1,2-benzoquinones. Adv. Heterocycl. Chem., 100, 1–51.

14 Horspool, W.M. (1969) Synthetic

15

16

17

18

19

20

21

22

23

24

1,2-quinones: synthesis and thermal reactions. Quart. Rev., 23, 204–235. Dürckheimer, W. and Paulus, E.F. (1985) Mechanism of purpurogallin formation: an adduct from 3-hydroxy-obenzoquinone and 4,5 dimethyl-obenzoquinone. Angew. Chem. Int. Ed. Engl., 24, 224–225. Musso, H., Figge, K., and Becker, D.J. (1961) Hydriergeschwindigkeit und Redoxpotential bei Chinonen. Chem. Ber., 94, 1107–1115. Clark, W.M. (1960) Oxidation–Reduction Potentials in Organic Systems, Williams & Wilkins, Baltimore, MD. Jovanovic, S.V., Kónya, K., and Scaiano, J.C. (1995) Redox reactions of 3,5-di-tertbutyl-1,2-benzoquinone. Implications for reversal of paper yellowing. Can. J. Chem., 73, 1803–1810. Namazian, M., Siahrostami, S., Noorbala, M.R., and Coote, M.L. (2006) Calculation of two-electron reduction potentials for some quinone derivatives in aqueous solution using Møller–Plesset perturbation theory. J. Mol. Struct., 759, 245–247. Kirk–Othmer Encyclopaedia of Chemical Technology (1968), vol. 16, 2nd edn, Interscience, New York, p. 899. Prota, G. (1992) Melanins and Melanogenesis, Academic Press, San Diego, CA. Ito, S. and Wakamatsu, K. (2006) Chemistry of melanins, in The Pigmentary System: Physiology and Pathophysiology, 2nd edn (eds J.J. Nordlund, R.E. Boissy, V.J. Hearing, R.A. King, W.S. Oetting, and J.-P. Ortonne), Blackwell, Malden, MA, pp. 282–310. Clews, J., Cooksey, C.J., Garratt, P.J., Land, E.J., Ramsden, C.A., and Riley, P.A. (2000) Oxidative cyclisation of N,N-dialkylcatecholamines to heterocyclic betaines via ortho-quinones: synthetic, pulse radiolytic and enzyme studies. J. Chem. Soc. Perkin Trans., 1, 4306–4315. Cooksey, C.J., Garratt, P.J., Land, E.J., Pavel, S., Ramsden, C.A., Riley, P.A., and

83

84

3 Biological Chemistry of o-Quinones

25

26

27

28

29

30

31

32

33 34

35

Smit, N.P.M. (1997) Evidence of the indirect formation of the catecholic intermediate substrate responsible for the autoactivation kinetics of tyrosinase. J. Biol. Chem., 272, 26226–26235. Land, E.J., Ramsden, C.A., and Riley, P.A. (2003) Tyrosinase autoactivation and the chemistry of ortho-quinone amines. Acc. Chem. Res., 36, 300–308. Ito, S. and Prota, G. (1977) A facile one-step synthesis of cysteinyldopas using mushroom tyrosinase. Experientia, 33, 1118–1119. Ito, S., Palumbo, A., and Prota, G. (1985) Tyrosinase-catalyzed conjugation of dopa with glutathione. Experientia, 41, 960–961. d’Ischia, M., Napolitano, A., Pezzella, A., Land, E.J., Ramsden, C.A., and Riley, P.A. (2005) 5,6-Dihydroxyindoles and indole-5,6-diones. Adv. Heterocycl. Chem., 89, 23–30. Land, E.J., Ramsden, C.A., and Riley, P.A. (2006) An MO study of regioselective amine addition to ortho-quinones relevant to melanogenesis. Tetrahedron, 62, 4884–4891. Land, E.J., Ramsden, C.A., Riley, P.A., and Yoganathan, G. (2003) Mechanistic studies of catechol generation from secondary quinone amines relevant to indole formation and tyrosinase activation. Pigment Cell Res., 16, 397–406. Land, E.J., Ramsden, C.A., and Riley, P.A. (2007) ortho-Quinone amines and derivatives: the inﬂuence of structure on the rates and modes of intramolecular reaction. Arkivoc, 2007 (xi), 23–36. Phillips, R.S., Fletcher, J.G., Von Tersch, R.L., and Kirk, K.L. (1990) Oxygenation of ﬂuorinated tyrosines by mushroom tyrosinase releases ﬂuoride ion. Arch. Biochem. Biophys., 276, 65–69. Raper, H.S. (1928) The aerobic oxidases. Physiol. Rev., 8, 245–282. Mason, H.S. (1948) The chemistry of melanin. Mechanism of the oxidation of dihydroxyphenylalanine by tyrosinase. J. Biol. Chem., 172, 83–92. Rorsman, H., Agrup, G., Hansson, C., and Rosengren, E. (1983) Biochemical recorders of malignant melanoma, in Malignant Melanoma: Advances of A

36

37

38

39

40

41

42

43

44

45

46

Decade (Pigment Cell 6) (ed. R.M. Mackie), Karger, Basel, pp. 93–115. Keene, J.P. (1964) Pulse radiolysis apparatus. J. Sci. Instrum., 41, 493–496. Land, E.J., Ito, S., Wakamatsu, K., and Riley, P.A. (2003) Rate constants for the ﬁrst two chemical steps of eumelanogenesis. Pigment Cell Res., 16, 487–493. Thompson, A., Land, E.J., Chedekel, M.R., Subbarao, K.V., and Truscott, T.G. (1985) A pulse radiolysis investigation of the oxidation of the melanin precursors 3,4-dihydroxyphenylalanine (dopa) and the cysteinyldopas. Biochim. Biophys. Acta, 843, 49–57. Land, E.J. and Riley, P.A. (2000) Spontaneous redox reactions of dopaquinone and the balance between the eumelanic and phaeomelanic pathways. Pigment Cell Res., 13, 273–277. Napolitano, A., Di Donato, P., Prota, G., and Land, E.J. (1999) Transient quinonimines and 1,4-benzothiazines of pheomelanogenesis: new pulse radiolytic and spectrophotometric evidence. Free Radic. Biol. Med., 27, 521–528. Edge, R., d’Ischia, M., Land, E.J., Napolitano, A., Navaratnam, S., Panzella, L., Pezzella, A., Ramsden, C.A., and Riley, P.A. (2006) Dopaquinone redox exchange with dihydroxyindole and dihydroxyindole carboxylic acid. Pigment Cell Res., 19, 443–450. Mason, H.S., Fowlks, W.L., and Peterson, E. (1955) Oxygen transfer and electron transport by the phenolase complex. J. Am. Chem. Soc., 77, 2914–2915. Pomerantz, S.H. (1966) The tyrosine hydroxylase activity of mammalian tyrosinase. J. Biol. Chem., 241, 161–168. Mason, H.S. (1955) Comparative biochemistry of the phenolase complex, in Advances in Enzymology, vol. 16 (ed. F.F. Nord), Interscience, New York, pp. 105–184. Lerner, A.B., Fitzpatrick, T.B., Calkins, E., and Summerson, W.H. (1949) Mammalian tyrosinase: preparation and properties. J. Biol. Chem., 178, 185–195. Lerch, K. (1981) Copper monooxygenases: tyrosinase and dopamine-betahydroxylase, in Metal Ions in Biological

References

47

48

49

50

51

52

53

54

55

56

57

Systems, vol. 13 (ed. H. Sigel), Decker, New York, pp. 143–186. Nelson, J.M. and Dawson, C.R. (1944) Tyrosinase, in Advances in Enzymology, vol. 4 (eds F.F. Nord and C.H. Werkman), Interscience, New York, pp. 99–152. Asimov, I. and Dawson, C.R. (1950) On the reaction inactivation of tyrosinase during aerobic oxidation of catechol. J. Am. Chem. Soc., 72, 820–828. Ingraham, L.L., Corse, J., and Makower, B. (1952) Enzymatic browning of fruits. III. Kinetics of the reaction inactivation of polyphenoloxidase. J. Am. Chem. Soc., 74, 2623–2626. Seiji, M., Sasaki, M., and Tomita, Y. (1978) Nature of tyrosinase inactivation in melanosomes. Tohoku J. Exp. Med., 125, 233–245. Tomita, Y., Hariu, A., Mizuno, C., and Seiji, M. (1980) Inactivation of tyrosinase by dopa. J. Invest. Dermatol., 75, 379–382. García-Cánovas, F., Tudela, J., MartínezMadrid, C., Varón, R., Garcia-Carmona, F., and Lozano, J.A. (1987) Kinetic study on the suicide inactivation of tyrosinase induced by catechol. Biochim. Biophys. Acta, 912, 417–423. Tudela, J., Garcia-Cánovas, F., Varón, R., Jiménez, M., Garcia-Carmona, F., and Lozano, J.A. (1988) Kinetic study in the transient phase of the suicide inactivation of frog epidermis tyrosinase. Biophys. Chem., 30, 303–310. Dietler, C. and Lerch, K. (1982) Reaction inactivation of tyrosinase, in Oxidases and Related Redox Systems (eds T.E. King, H.S. Mason, and M. Morrison), Pergamon, New York, pp. 305–317. Land, E.J., Ramsden, C.A., and Riley, P.A. (2007) The mechanism of suicideinactivation of tyrosinase: a substrate structure investigation. Tohoku J. Exp. Med., 212, 341–348. Land, E.J., Ramsden, C.A., Riley, P.A., and Stratford, M.R.L. (2008) Evidence consistent with the requirement of cresolase activity for suicide inactivation of tyrosinase. Tohoku J. Exp. Med., 216, 231–238. Ramsden, C.A., Stratford, M.R.L., and Riley, P.A. (2009) The inﬂuence of catechol structure on the suicide

58

59

60

61

62

63

64

65

66

67

inactivation of tyrosinase. Org. Biomol. Chem., 7, 3388–3390. Spritz, R.A., Ho, L., Furumara, M., and Hearing, V.J. (1997) Mutational analysis of copper binding by human tyrosinase. J. Invest. Dermatol., 109, 207–212. Muñoz-Muñoz, J.L., Garcia-Molina, F., Garcia-Ruiz, P.A., Molina-Alcaron, M., Tudela, J., Garcia-Cánovas, F., and Rodriguez-Lopez, J.N. (2008) Phenolic substrates and suicide inactivation of tyrosinase: kinetics and mechanism. Biochem. J., 416, 431–440. Ramsden, C.A. and Riley, P.A. (2010) Mechanistic studies of tyrosinase suicide inactivation. Arkivoc, 2010 (i), 260–274. Eicken, C., Zippel, F., BüldtKarentzopoulos, K., and Krebs, B. (1998) Biochemical and spectroscopic characterization of catechol oxidase from sweet potatoes (Ipomoea batatas) containing a type-3 dicopper center. FEBS Lett., 436, 293–299. Gerdemann, C., Eicken, C., and Krebs, B. (2002) The crystal structure of catechol oxidase: new insight into the function of type-3 copper proteins. Acc. Chem. Res., 35, 183–191. Klabunde, T., Eicken, C., Sacchettini, J.C., and Krebs, B. (1998) Crystal structure of a plant catechol oxidase containing a dicopper center. Nat. Struct. Biol., 5, 1084–1090. Matoba, Y., Kumagai, T., Yamamoto, A., Yoshitsu, H., and Sugiyama, M. (2006) Crystallographic evidence that dinuclear copper center of tyrosinase is ﬂexible during catalysis. J. Biol. Chem., 281, 8981–8990. Decker, H., Schweikardt, T., and Tuczek, F. (2006) The ﬁrst crystal structure of tyrosinase: all questions answered? Angew. Chem. Int. Ed., 45, 4546–4550. Van Gastel, M., Bubacco, L., Groenen, E.J.J., Vijgenboom, E., and Canters, G.W. (2000) EPR study of the dinuclear active copper site of tyrosinase from Streptomyces antibioticus. FEBS Lett., 474, 228–232. Bubacco, L., van Gastel, M., Groenen, E.J.J., Vijgenboom, E., and Canters, G.W. (2003) Spectroscopic characterization of the electronic changes in the active site of Streptomyces antibioticus tyrosinase upon

85

86

3 Biological Chemistry of o-Quinones binding of transition state analogue inhibitors. J. Biol. Chem., 278, 7381–7389. 68 Ramsden, C.A. and Riley, P.A. (2010) Studies of the competing rates of catechol oxidation and suicide inactivation of tyrosinase. Arkivoc, 2010 (x), 248–255. 69 Hansson, C., Rorsman, H., and Rosengren, E. (1980) 5-Hydroxydopa, a new compound in the Raper–Mason scheme of melanogenesis. Acta Derm. Venereol., 60, 281–186. 70 Agrup, G., Rorsman, H., and Rosengren, E. (1982) 5-OH-dopa, product of and

substrate for tyrosinase. Acta Derm. Venereol., 62, 371–376. 71 Carlberg, M., Jergil, B., Lindbladh, C., and Rosengren, E. (1984) Enzymatic 5-hydroxylation of l-dopa by a tyrosinase isolated from the sea anemone Metrium senile. Gen. Pharmacol., 15, 301–307. 72 Burzio, L.A. and Waite, J.H. (2002) The other Topa: formation of 3,4,5-trihydroxyphenylalanine in peptides. Anal. Biochem., 306, 108–114.

87

4 Biosynthesis of Melanins José Carlos García-Borrón and M. Concepción Olivares Sánchez

4.1 Introduction

Melanogenesis, the biochemical pathway responsible for the biosynthesis of the chemo- and photoprotective melanin pigments mostly from phenolic precursors, is a widespread process occurring in all types of organisms throughout the phylogenetic scale. In prokaryotes, melanogenesis consists of a rather simple series of reactions involving just one melanogenic enzyme, while in mammals it is a tightly regulated pathway coordinated by the combined action of a family of melanocytespeciﬁc gene products, some of which associate to form a multienzymatic complex. The mammalian melanogenic enzymes are three related and highly similar metalloproteins: tyrosinase, tyrosinase-related protein 1 (Tyrp1 or gp75) and tyrosinaserelated protein 2 (Tyrp2, also called dopachrome tautomerase (Dct), and designated here Dct/Tyrp2). These proteins share numerous structural similarities, and follow quite similar biosynthetic, processing, and trafﬁcking pathways [1]. All three enzymes are type 1 membrane-bound melanosomal glycoproteins. They are synthesized by ribosomes and transported through the rough endoplasmic reticulum (ER) and the Golgi apparatus, where they undergo post-translational processing and glycosylation. They are then delivered to the early melanosomes and the synthesis of melanin ensues. The present chapter describes the currently accepted biochemical pathway for melanin synthesis, which is in fact a modiﬁed version of the Raper–Mason route outlined in the ﬁrst half of the twentieth century. We summarize the current knowledge of the structural features and catalytic mechanism of the melanogenic enzymes, focusing on the mammalian proteins (mainly tyrosinase and Dct/Tyrp2, which are better characterized). We do not address speciﬁc aspects of the melanogenic pathway in prokaryotes or in plants, and the reader interested in these organisms is referred to one of various recent and authoritative reviews [2–5]. We also outline several aspects of the regulation of the melanogenic pathway in order to give an integrated view of mammalian melanogenesis. We also refer to other chapters of this book for further details on related topics such as the genetics of the melanogenic proteins, the biogenesis of the melanosome, and the chemical Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

88

4 Biosynthesis of Melanins

structure of the melanin pigment, as they are deeply related to those covered here, but beyond the scope of the present chapter.

4.2 Raper–Mason Pathway

In vertebrates, melanins are formed from the amino acid l-tyrosine by means of a unique series of enzymatic and chemical reactions known as the Raper–Mason pathway. Based on the observation that melanins could be formed from phenolic precursors in the presence of puriﬁed tyrosinase, the melanogenic pathway was originally delineated by Raper [6] and Mason [7] taking into account the involvement of a single enzyme, tyrosinase, as well as a series of spontaneous chemical reactions. However, comparison of the structure of natural melanins and melanins synthesized in vitro by the action of tyrosinase on several melanogenic precursors, as well as the study of the action of melanosomal extracts on melanin biosynthesis intermediates, led to the discovery of several other enzymatic melanogenic activities in addition to tyrosinase. These enzymatic activities have been incorporated to the currently accepted version of the Raper–Mason pathway. In addition, genetic evidence arising from the study of mutant mice with various coat color phenotypes demonstrates that certain nonenzymatic melanosomal proteins such as Pmel-17, the product of the silver locus [8], act to support a correct melanogenic function. For simplicity, the Raper–Mason pathway can be divided into two phases. A proximal phase contains the rate-limiting tyrosinase-catalyzed reactions and leads to the formation of a key intermediate, l-dopachrome, from the amino acid ltyrosine. In the subsequent distal phase, l-dopachrome evolves spontaneously and/or enzymatically to reactive intermediates and the chemical polymerization steps that ultimately yield the melanogenic pigments take place. 4.2.1 Phase I Melanogenesis: The Proximal Raper–Mason Pathway – From L-tyrosine to L-dopachrome

The key regulatory and rate-limiting melanogenic enzyme is tyrosinase, a coppercontaining, membrane-bound glycoprotein located in the melanosomes. This enzyme is able to catalyze the ﬁrst two steps of the proximal phase: the hydroxylation of l-tyrosine and the subsequent oxidation of the intermediate o-diphenol (l-3,4-dihydroxyphenylalanine (l-dopa)) to yield l-dopaquinone (monophenolase or cresolase activity, EC 1.14.18.1, and diphenolase or catechol oxidase activity, EC 1.10.3.1, respectively) (Figure 4.1). Hydroxylation of l-tyrosine is the rate-limiting step in melanin synthesis. Although tyrosine hydroxylase and dopa oxidase activities occur in the same active site [10], the diphenolase activity is approximately 7 times faster than monophenolase activity. The mechanism of these reactions has been controversial and will be discussed below (Section 4.3.2). In any case, there is no doubt that the ﬁnal product of tyrosinase action on l-tyrosine, l-dopaquinone,

4.2 Raper–Mason Pathway

The mammalian melanogenic pathway. Tyrosinase is the only enzyme in the proximal phase of melanogenesis. The role of the Tyrps in the distal phase is critical to determine the type and amount of eumelanins synthesized: l-dopachrome can undergo a spontaneous decarboxylative

Figure 4.1

reaction to form DHI or a nondecarboxylative rearrangement catalyzed by Dct/Tyrp2 and subsequent oxidation to IQCA catalyzed by Tyrp1 (at least in mouse melanocytes). The pathway for pheomelanin synthesis is less well understood. (Adapted from [9].)

is the ﬁrst branch point of two connected biosynthetic routes in melanogenesis leading to the two main types of melanin pigments: dark (brown/black) eumelanin or light (yellow/red) pheomelanin [1]. During eumelanin biosynthesis (Figure 4.1, left), l-dopaquinone can undergo two types of spontaneous chemical reactions: an intramolecular 1,4-addition to the benzene ring and/or a water addition reaction. In the ﬁrst case, the amino group of the l-dopaquinone side-chain engages in an intramolecular Michael addition with cyclization to yield l-cyclodopa (also termed leucodopachrome). This intermediate is quickly and spontaneously oxidized to l-dopachrome by another molecule of l-dopaquinone, which is reduced back to l-dopa [11, 12]. Both reactions are fast and proceed without enzymatic control, although cyclization of ldopaquinone is favored over conjugation with thiols and other nucleophilic chemicals due to its intramolecular nature and to the presumably low concentrations of these compounds inside the eumelanosome. Note that half the l-dopa oxidized by tyrosinase to l-dopaquinone is reduced back to l-dopa. This is most likely a relevant feature of the pathway owing to the regulatory role of l-dopa on the tyrosine hydroxylase activity of the enzyme (see below). On the other hand, the formation of l-cyclodopa is favored as the pH increases, because the amino group should be

89

90

4 Biosynthesis of Melanins

nonprotonated to allow for the nucleophilic attack on position 6. Therefore, at acidic intramelanosomal pH values cyclization should be rather slow and water addition to the benzene ring may become possible, thus leading to the formation of a highly reactive three-hydroxylated phenol, topa (3,4,5-trihydroxyphenylalanine). Presumably, topa would also evolve to l-dopachrome through a series of slow spontaneous chemical reactions ([13]; not shown in Figure 4.1). 4.2.2 Distal Melanogenic Steps: From L-Dopachrome to Eumelanins

l-Dopachrome is considered the ﬁnal product of the proximal phase in the Raper– Mason pathway and is in fact a second branch point in melanogenesis leading to slightly different types of eumelanins, with varying proportions of carboxylated or decarboxylated monomeric units. The distal melanogenic reactions begin with the spontaneous, nonenzymatic decarboxylative rearrangement of l-dopachrome to 5,6-dihydroxyindole (DHI) and CO2. DHI is then oxidized to 5,6-indolequinone (IQ). Under aerobic conditions, this oxidation occurs spontaneously, but tyrosinase signiﬁcantly accelerates the rate of the reaction. Alternatively, orange red l-dopachrome may be enzymatically transformed into colorless DHI-2-carboxylic acid (DHICA) [14]. The corresponding catalytic activity was originally designated dopachrome conversion factor, then dopachrome isomerase, and is now named Dct (EC 5.3.3.12), also known as Tyrp2 [15]. It is worth noting that traces of divalent transition metal cations, such as Cu2+ or Ni2+, also bring about this isomeric rearrangement of l-dopachrome, and can yield mixtures of DHI and DHICA [16]. Although the enzymatic and chemical conversions of l-dopachrome are not mutually exclusive, the enzymatically catalyzed reaction is highly stereo-speciﬁc for l-dopachrome and, inside the melanosome, the level of Dct activity controls the DHICA/DHI ratio [17]. DHICA is relatively stable at physiological pH and temperature, and does not lose its carboxyl group easily to form DHI. Further oxidation of DHICA yields indole-2-carboxylic acid-5,6-quinone (IQCA). Spontaneous oxidation is much slower for DHICA than for DHI because of the electron-withdrawing effect of the carboxyl group at position 2, so that DHICA oxidation in vivo is most likely an essentially enzymatic reaction. However, the details of the enzymology of DHICA oxidation are complex and have not yet been completely worked out. Indeed, it has been shown that whereas mouse tyrosinase is unable to recognize DHICA as a diphenolic substrate to promote its oxidation, mouse Tyrp1 behaves as a low-speciﬁc-activity DHICA oxidase [18, 19]. Conversely, human tyrosinase shows DHICA oxidase activity [20]. Therefore, it is possible that the oxidation of DHICA in vivo is carried out by different enzymes, Tyrp1 or tyrosinase, depending on the particular mammalian species considered. Accordingly, the physiological role of Tyrp1 in some species, especially in human melanocytes, has been and still is a matter of discussion (see below). In any case, it is ﬁrmly established that l-dopachrome formed in the initial phase of the melanogenic pathway is transformed into mixtures of DHICA and DHI,

4.2 Raper–Mason Pathway

and that the relative proportions of these two diphenols are dictated by a complex set of intramelanosomal conditions with a major role for Dct/Tyrp2 activity. The next steps in eumelanogenesis are less well characterized. DHI, DHICA, and their corresponding o-quinones undergo relatively unordered and intermixed spontaneous polymerization to form black, insoluble eumelanin (DHI-melanin) and golden-brown, poorly soluble eumelanin (DHICA-melanin). The size, chelating, and absorption properties of the ﬁnal eumelanin depend on the initial DHICA/ DHI ratio [14, 17]. DHICA-melanin exerts a stronger antioxidant effect than DHImelanin, and this behavior may be due to the different structure and solubility of the two melanins. Furthermore, the DHICA branch of eumelanogenesis is less cytotoxic than the DHI branch. In fact, the oxidative metabolism of DHI, and to a lesser extent DHICA, is a source of cytotoxic species, including highly reactive quinone intermediates and oxygen species such as hydrogen peroxide [21]. Tyrp1 and Tyrp2 are thought to be critical to protect melanocytes from the inherent toxicity of melanogenic reactions. Indeed, in the presence of these proteins the synthesis of DHI is kept to a minimum, whereas the production and oxidative metabolism of DHICA is favored, so that melanogenesis follows primarily the less-toxic DHICA pathway. Given our incomplete knowledge of the distal portion of melanogenesis, one open question is the possible involvement of other melanosomal proteins different from tyrosinase and the Tyrps in the control of the distal steps of the pathway. In this respect, the contribution of the silver protein as an accelerating factor for the polymerization of monomeric intermediates has been proposed [22, 23], but a precise catalytic activity has never been deﬁnitively proven for this protein (discussed below). In summary, the type of melanin produced depends not only on the availability of substrates [24], but also on the relative activities of at least three melanogenic enzymes, which leads to a reﬁned control of melanosynthesis in animals compared with lower organisms [17] and accounts for the occurrence of a wide spectrum of natural melanin pigments with subtly different physicochemical properties. 4.2.3 Biosynthesis of Pheomelanins

The switch from eumelanin biosynthesis to production of pheomelanins depends upon the availability of low-molecular-weight sulfydryl compounds such as the amino acid l-Cys or reduced glutathione. In the presence of these compounds, the rapid conjugation between l-dopaquinone and the thiol group yields a family of sulfur-containing intermediates related to pheomelanins [25, 26]. However, the pheomelanogenic intermediates and the regulatory events that control the availability of thiolic substrates within melanosomes are poorly known. All the reactions in the pheomelanin-speciﬁc pathway are thought to involve highly unstable intermediates and, accordingly, they might occur spontaneously. In fact, efforts to identify melanosomal enzymes responsible for the catalysis or regulation of these

91

92

4 Biosynthesis of Melanins

reactions have been largely unsuccessful. A simple model of pheomelanogenesis (Figure 4.1, right) assumes that the nucleophilic attack of the thiol group of l-Cys to l-dopaquinone yields mainly 5-cysteinyldopa [27]. However, in addition to carbon 5 other free positions in the l-dopaquinone ring such as 2 and 6 are also reactive, so that a complex mixture of isomers is obtained. 5-cysteinyldopa would undergo several structural rearrangements and dehydration to form an alanylhydroxy-benzothiazine monomer, the proposed monomeric subunit for pheomelanin. However, pheomelanin biosynthesis could proceed through a more complex pathway involving glutathione rather than Cys, since this tripeptide is much more abundant in the intracellular medium than the free amino acid. Accordingly, it has been proposed that the ﬁrst major thiol-conjugated species formed inside melanocytes is 5-glutathionyldopa. A dipeptidase would then catalyze the release of 5-cysteinyldopa and the pathway would progress as described above ([28]; not shown in Figure 4.1 for simplicity). This glutathione-dependent branch may still be an oversimpliﬁcation of the in vivo situation, where still uncharacterized enzymatic activities might be involved.

4.3 Structural and Functional Properties of the Melanogenic Enzymes

As discussed above, tyrosinase is the key melanogenic regulatory enzyme in vertebrates. In plants, the highly similar catechol oxidases catalyze the oxidation of o-diphenols, but they are unable to carry out the hydroxylation of monophenolic substrates. The active sites of the two types of enzymes (termed phenoloxidases) are very similar in terms of their sequence and physicochemical properties. Phenoloxidases and the oxygen carrier proteins hemocyanins belong to the type 3 copper protein family. Collectively, they are responsible for pigmentation of skin and hair, browning of fruits, and wound healing in plants and arthropods. Since melanin pigments are found in all types of organisms, tyrosinases are ubiquitously distributed throughout the phylogenetic scale, but the following discussion is focused on the mammalian enzymes. 4.3.1 Structure of Tyrosinase and Related Proteins

The overall structure and the active site of tyrosinases are highly conserved among different vertebrate species. Moreover, mammalian tyrosinases show high homology with Tyrp1 and Dct/Tyrp2 (up to 40% amino acid identity and approximately 70% amino acid homology [29]). There are several residues strictly conserved in all tyrosinases, the more important being the copper-binding ligands and other amino acids establishing interactions critical to maintain the globular folding of the protein. In addition, tyrosinases from higher organisms display some other conserved structural features, such as the C-terminal motifs for trafﬁcking and targeting, the Cys clusters, several N-glycosylation sites, and a putative transmem-

4.3 Structural and Functional Properties of the Melanogenic Enzymes a)

b)

(a) Schematic representation of the tyrosinase family proteins. The numbering, precise positions of domains, and amino acid sequence of metal binding sites correspond to the mouse proteins. SP, signal peptide; Cys, cysteine-rich segments; Cu or Me, copper- or metal-binding domains (sequence comprised between the ﬁrst and third His detailed below); TM, transmembrane fragment. (b) Proposed consensus for tyrosinases [30]. Bold: the six His directly bound to Cu. Subscript indicates the ﬁrst (A)

Figure 4.2

or second (B) binding site and the order of the His in the primary sequence. Φ means aromatic residue (F, Y, or W). B means hydrophobic residue. The residues are labeled with a subscript indicating their position in relation to the closest H. Residues in italics mean that the residue changes from tyrosinase to Tyrps. Underlined: conserved residues important to maintain the three-dimensional structure of tyrosinase and its family.

brane domain near its C-terminus that ensures their association with the melanosomal membrane in a similar orientation relative to the melanosomal lumen [30] (Figure 4.2). Most of these elements are also conserved in mammalian Tyrps. However, in spite of the remarkable sequence similarity of their active site, the three proteins of the tyrosinase family have distinct enzyme activities and function at different steps of melanin biosynthesis [1]. This is most likely a consequence of the speciﬁc binding of different metal cofactors in the two conserved metal-binding motifs that are critical to their enzymatic functions: copper in tyrosinase [31] and zinc in Dct/Tyrp2 [32]. The precise metal cofactor in Tyrp1 is still unknown [33]. So far, no mammalian tyrosinase has been crystallized, probably because it is a somewhat heterogeneous transmembrane glycosylated protein. Nevertheless, a variety of data based on mutagenesis studies [34, 35], the available crystallographic data on a plant catechol oxidase (Ipomoea batatas) [36], and the ﬁrst published crystal structure of a prokaryotic tyrosinase (Streptomyces castaneoglobisporus) [37]

93

94

4 Biosynthesis of Melanins

allow a model for the mammalian tyrosinase active site and reaction mechanism to be deﬁned. In this model, the two copper-binding sites in tyrosinase, named CuA and CuB, are peptidic regions containing three perfectly conserved His residues. CuA displays a H-xn-H-x8-H motif and CuB displays a H-x3-H-xn-H motif, where “n” is a variable number of residues [30]. The six ion-binding His residues are adjacent due to the folding of the protein at the active site, consisting of a hydrophobic pocket inside a helix bundle with four densely packed helices. The overall structure is maintained by electrostatic and cation-π interactions. Oxygen binds as a side-on (μ–η2:η2) peroxide bridge [38] and, in the resting form of the enzyme, the pair of antiferromagnetically coupled copper ions are pentacoordinated in a distorted square pyramidal geometry [39]. Although it is generally accepted that each copper ion is bound only by the three His residues in either CuA and CuB, some other alternative modes of cofactor quelation have been proposed, with four putative ligands (one Cys and three His) [40]. Apart from the invariant His residues in the copper-binding sites, all tyrosinases throughout nature display several other strictly conserved residues most likely involved either in the binding and docking of the substrates or in the maintenance of the structural integrity of the active site. Accordingly, we proposed a broader consensus for the copper-binding regions with a new notation to distinguish residues essential for tyrosinase active-site folding [30] (Figure 4.2). This allows for the correlation of the available crystallographic data and of studies on the functional effects of naturally occurring or artiﬁcially created mutations, with the contribution of these residues to the active-site structure or their potential catalytic role [35, 41, 42]. Notably, some of these residues are also present in the Tyrps. The structure of all mammalian tyrosinases characterized to date also comprises the following common features [30] (Figure 4.2). The N-terminal signal peptide targets the nascent polypeptides to the ER for its folding, processing, and sorting. The signal peptide must be cleaved for correct folding due to its hydrophobicity. It is generally assumed that cleavage takes place between G18 and H19 in human and mouse tyrosinase. Within the carboxyl tail of the protein, mammalian tyrosinases contain a single transmembrane segment followed by a short extension. There is no doubt that the bulk of the protein containing the active site faces the lumen of the melanosome, whereas the short C-terminal extension is located in the cytosol. The transmembrane fragment anchors the protein to the melanosomal membrane. The C-terminal cytosolic peptide contains essential signals for targeting the enzyme to the melanosome [43, 44]. Some of these signals are a dileucine motif (LL, with some variants) and the tyrosine-based motif (YXXB, where B is any bulky hydrophobic residue) [45]. Mammalian tyrosinase family proteins also display conserved Cys-rich domains. The number and relevance of Cys residues in tyrosinase is variable. The Streptomyces enzyme is completely devoid of Cys, while mammalian tyrosinases have 17 Cys residues clustered in three regions. Notably, the second cluster has been denominated the epidermal growth factor (EGF)-like region [46] as it partially matches the two EGF-like motifs and an EGFrelated domain, the laminin-LE. Plant catechol oxidases contain only the ﬁrst cluster. Even though almost all Cys seem to be important for a correct protein

4.3 Structural and Functional Properties of the Melanogenic Enzymes

folding and copper acquisition [47, 48], the function of the clusters and the location of disulﬁde bridges in mammalian tyrosinases and Tyrps remain mostly unknown. In higher organisms, the tyrosinase family proteins are glycoproteins containing several N-glycosylation sites. The role of N-glycans in sorting, stability, and activity of mammalian tyrosinase has been extensively studied. Post-translational processing of tyrosinase and its trafﬁc through the secretory pathway are critical for the acquisition of an active form of the enzyme. Some N-glycosylation sequons are strongly conserved, such as the one present within the CuB site. This sequon appears particularly important for the correct folding of the enzyme [49]. When initially synthesized, the nonglycosytaled protein backbone of tyrosinase has an apparent molecular weight of 55 kDa; after terminal glycosylation, the electrophoretic pattern of tyrosinase is indicative of some heterogeneity and the molecular weight of the mature protein shifts to about 65–75 kDa [50]. Tyrosinase folding and maturation is assisted by ER-resident chaperones, with the interaction of calnexin and the carbohydrate chains in tyrosinase playing an important role [49, 51]. Although it is still unclear exactly where or how copper ions are transferred into apo-tyrosinase, it is accepted that terminal glycosylation and copper acquisition of mammalian tyrosinase occurs as the enzyme is processed through the Golgi complex and delivered to stage II melanosomes via early endosomal intermediates [44, 52], where enzyme activity is expressed. In fact, it has been proposed that late copper binding could prevent abnormal and undesirable melanogenesis in proximal compartments of the biosynthetic-secretory pathway [53]. Moreover, the involvement of the Menkes copper transporter (MNK), located in the trans-Golgi network, has been demonstrated [54]. Interestingly, it has been shown that proper tyrosinase folding and post-translational glycosylation can occur in the absence of copper loading [49]. This suggests that copper is transferred to a well-structured active site able to bind the metal ion with high afﬁnity, thus avoiding leakage of the cofactor. Since tyrosinase is the key and rate-limiting melanogenic enzyme, loss-offunction mutations on the tyrosinase gene result in reduction or absence of melanin of melanocytes in the skin, hair follicles, and eyes, causing oculocutaneous albinism (OCA) type 1 (tyrosinase-negative, OMIM 203100) [55]. As discussed below, OCA1 is often associated with retention of defective tyrosinase in the ER and subsequent degradation [56]. At present, more than 200 mutations in the tyrosinase gene have been associated with albinism (reviewed by Oetting [57]). Tyrosinase and the Tyrps exist as a high-molecular-weight complex with relatively stable interactions putatively involving the highly conserved Cys-rich EGF domains. The strong intermolecular association of Tyrp1 and tyrosinase enhances the intracellular stability of tyrosinase [58, 59] and, as discussed below, has a signiﬁcant effect on its maturation and intracellular transport [60]. 4.3.2 Catalytic Cycle of Tyrosinase

The current view of the mechanism of tyrosinase catalysis considers several forms of the enzyme and two connected cycles for the cresolase (tyrosine hydroxylase)

95

96

4 Biosynthesis of Melanins

and catecholase (dopa oxidase) activities, occurring in a single active site. Depending on the absence/presence of oxygen and the oxidation state of the copper ions [Cu(II)/Cu(I)], the enzyme can exist in three interconvertible forms, called met, oxy, and deoxy. The mechanism is well supported by structural data from the ﬁrst published crystal structure of a tyrosinase (from S. castaneoglobisporus) [37]. This structure provides a good model to interpret the function of residues in the active site of the mammalian tyrosinases due to the high degree of conservation within this region in all known tyrosinases. Indeed, homology modeling of mouse tyrosinase based on this structure already provided a framework to successfully explain the effects on enzyme kinetics of artiﬁcial and naturally occurring albino mutations [61]. Both the tyrosine hydroxylase or dopa oxidase cycles are initiated when the enzyme is in the oxy state and the appropriate substrates enter the active site (Figure 4.3). However, there are important differences between the two types of reactions. For instance, the tyrosine hydroxylase activity, but not the dopa oxidase activity, shows a characteristic lag period before the reaction reaches its maximal rate. This lag phase can be shortened by catalytic amounts of l-dopa [63]. Therefore, l-dopa is not only the product of the monophenolase activity of tyrosinase acting on l-tyrosine as substrate and a substrate for the catecholase activity, but also a cofactor for the tyrosine hydroxylase cycle. Moreover, the afﬁnity of mammalian tyrosinases for l-dopa acting as cofactor for tyrosine hydroxylase activity is about two orders of magnitude higher than those for the compound acting as a catecholic substrate [64]. The mechanism that follows accounts for these kinetic features as well as for some other differences in the catalytic requirements of tyrosine hydroxylase and dopa oxidase activities. 4.3.2.1 Cresolase (Tyrosine Hydroxylase) Reaction Cycle In the oxy form, molecular oxygen is coordinated between the two active-site copper ions, each of which is bound to the protein matrix by three His residues held in place by a pair of antiparallel α-helices. The only exception is one His in the CuA site (His54 in the S. castaneoglobisporus enzyme and His202 in mouse tyrosinase), located in a ﬂexible loop. This residue could be responsible for a proton shift from the ortho position of l-tyrosine during the catalytic cycle of the Streptomyces enzyme [65]. The tyrosine hydroxylase cycle is initiated by the binding of l-tyrosine to the active site (Figure 4.3). The structures mentioned before provide information on the possible orientation of monophenols at the active site, although this aspect may not be universal. At least in mammalian tyrosinases, the most likely orientation of l-tyrosine would be forced by the π–π interaction between the aromatic ring in the substrate and the second His in the CuB site. One copper of the cluster forms a tyrosine hydroxylate complex placing carbon 3 of the substrate in position to accept the terminal oxygen of the peroxide, which is bound to the second copper, to form l-dopa and then l-dopaquinone. The o-quinone then leaves the reduced bicuprous site, allowing for the entrance of a new oxygen molecule. Therefore, even in the tyrosine hydroxylase cycle the released substrate should be l-dopaquinone [66] rather than l-dopa, as previously suggested. Oxygen

4.3 Structural and Functional Properties of the Melanogenic Enzymes

Mechanism of action for tyrosinase. The two catalytic cycles of tyrosinase are depicted (the His numbering corresponds to the mouse enzyme). Note that met-tyrosinase is involved in the dopa

Figure 4.3

oxidase (DO) cycle, but not in the tyrosine hydroxylase (TH) cycle: when l-tyrosine binds to this form of the enzyme, a dead-end complex is formed. (Adapted from [62].)

97

98

4 Biosynthesis of Melanins

binding then induces an oxidative change in the valence of the copper ions: the two oxygen atoms bind to the Cu(I)–Cu(I) center of deoxy-tyrosinase to yield a peroxide bound Cu(II)–Cu(II) center in oxy-tyrosinase. The unusual enzyme kinetics observed in this reaction are due to the nonenzymatic indirect formation of l-dopa by the chemical reactions accompanying dopaquinone cyclization [67] (Figure 4.1). On the other hand, plant and fungal catechol oxidases lack tyrosine hydroxylase activity despite the high similarity with animal tyrosinases. This could be explained by the presence in all catechol oxidases sequenced this far of a bulky aromatic residue shielding the CuA site [36], which is not found in animal tyrosinases. In keeping with this possibility, it has been proposed that monophenols would dock to CuA whereas o-diphenols would dock to CuB in the active site of mammalian tyrosinases [35]. 4.3.2.2 Catecholase (Dopa Oxidase) Reaction Cycle In the dopa oxidase cycle, the o-diphenolic substrate would bind to the oxy form of the enzyme. Binding of the catechol to CuB would also labilize the oxygen, resulting in the oxidation of the substrate and release of l-dopaquinone. Biochemical and kinetic evidences suggest that the His residue adjacent to the third copper ligand in the CuB site is involved in stereospeciﬁc binding of diphenols, but not monophenols [35], and that binding involves only one of the two available hydroxyl groups in the substrate. The met bicupric state of the enzyme accounts for the kinetic characteristics of the lag phase in the tyrosine hydroxylase cycle previously mentioned. This enzymatic form can bind either l-tyrosine or l-dopa. l-Dopa would dock to the two copper ions through both catecholic hydroxyl groups, hence with higher afﬁnity than when the binding proceeds only through CuB (oxy form). This differential mode of binding explains the 100-fold higher afﬁnity of l-dopa acting as a cofactor than as a substrate [64]. Moreover, this orientation accelerates the oxidation of the substrate and the reduced deoxy-tyrosinase is reoxidized upon oxygen binding. Docking of l-tyrosine to met-tyrosinase at the catalytic center would lead to a dead-end complex (Figure 4.3). The inhibition by high concentrations of l-tyrosine reported for tyrosinase [68] could be related to the formation of this dead-end complex. Finally, tyrosinase is characterized by an irreversible inactivation that occurs during the oxidation of catechols. A recently proposed mechanism accounting for this suicide inactivation is based on the ortho-hydroxylation of catecholic substrates, where Cu(II) is reduced to Cu(0), which no longer binds to the enzyme and is eliminated (reductive elimination) [69]. This process is dependent on the cresolase activity of tyrosinase and, accordingly, it is not exhibited by enzymes lacking tyrosine hydroxylase activity [70]. 4.3.3 Dct/Tyrp2

Dct/Tyrp2 is encoded at the slaty locus [71], whose mutation in mice is responsible for the slaty phenotype, characterized by the production of DHICA-poor eumela-

4.3 Structural and Functional Properties of the Melanogenic Enzymes

nin. Dct/Tyrp2 shows remarkable homology to tyrosinase and the protein displays most of the salient conserved structural elements discussed above. However, although Dct/Tyrp2 is also associated to the melanosomal membrane by a single membrane-spanning helix, the sorting and trafﬁcking signals in its short cytoplasmic C-terminal extension are different from those in tyrosinase, suggesting a differential intracellular processing [72]. Dct/Tyrp2 displays several glycosylation sequons, whose occupancy is important for stability and enzymatic function [73]. The sequences of the metal ion binding sites in Dct/Tyrp2 (MeA and MeB) share with tyrosinase the position and number of the His residues likely involved in chelation of the metal cofactor. However, the nature of the metal cofactor is different. Puriﬁed Dct/Tyrp2 contains bound zinc ions and enzymatic activity can be reconstituted from apoenzymatic preparations exclusively by addition of zinc, but not by copper or iron ions [32, 74]. Therefore, it is now widely accepted that Dct/ Tyrp2 is a zinc protein. This is consistent with the nature of the reaction catalyzed by Dct: Zn(II) is devoid of redox properties and therefore useless for oxidative reactions, but is highly efﬁcient as a tautomerization catalyst. Accordingly, the presence of zinc ions instead of copper in the metal-binding site is considered the most important factor determining the catalytic events at the active site of Dct/ Tyrp2. Each Zn(II) is probably bound to the protein moiety of the enzyme by the three His residues conserved in the MeA and MeB sites, most likely forming a distorted tetrahedron, the typical geometry of this ion. l-dopachrome would bind to the Zn(II) ions through the semiquinonic face of the molecule, displacing a water molecule bound to the fourth position of the tetrahedron, so that each ion would remain linked to the polypeptidic chain by the three His residues (Figure 4.4). This enzyme–substrate complex initiates an electronic rearrangement in the indolic ring with subsequent hydrogen migration from position 3 to position 2, leading to the formation of DHICA as the ﬁnal product of the reaction. Although the mouse and human enzymes act on l-dopachrome stereospeciﬁcally to release DHICA as the unique product, there are some reports on Dctrelated enzymes in mammals and lower organisms able to act on d-dopachrome or dopaminochrome to give DHI [75, 76]. The speciﬁcity of murine and human Dct/Tyrp2 for the l-enantiomer could be explained by the stereospeciﬁc interaction of the carboxyl group of l-dopachrome with some residue different from the conserved histidines. This amino acid would anchor the substrate at the active site with the appropriate orientation, similarly to tyrosinase with l-dopa [35]. Unfortunately, attempts to identify this residue have been unsuccessful [77]. In addition to its role in the metabolism of carboxylated melanogenic intermediates, Dct/Tyrp2 might also fulﬁll still uncharacterized functions. In keeping with this possibility, Dct/Tyrp2 is expressed earlier in the developing mouse than either tyrosinase or Tyrp1 and hence is already present in melanoblasts at developmental stages where no l-dopachrome is expected to be formed [78]. Moreover, a recent report revealed an intriguing link between Dct/Tyrp2 and hypoxia-inducible factor (HIF), a transcription factor that regulates cellular responses to changes in oxygen concentration and has an antiapoptotic effect against DNA-damage-induced apoptosis in certain model systems. This antiapoptotic activity of HIF-1 was shown to

99

100

4 Biosynthesis of Melanins

Figure 4.4 Mechanism of dopachrome tautomerization catalyzed by Dct/Tyrp2. In this catalytic cycle the enzyme presents only the met form and no redox reactions or oxygen binding take place at the active site.

l-Dopachrome binds to zinc ions through the semiquinonic part of the molecule, and subsequent electronic and proton rearrangements lead to DHICA. (Adapted from [62].)

depend on the transcriptional upregulation of Dct/Tyrp2 in a deﬁned population of sensory neurons in Caenorhabditis elegans. The TYR-2 homolog is secreted by ASJ sensory neurons to antagonize CEP-1-dependent germline apoptosis (CEP-1 is the homolog of the tumor suppressor p53). Importantly, the antiapoptotic effect of Dct/Tyrp2 might be a general feature of the enzyme and an evolutionarily conserved function, since knock-down of the protein in human melanoma cells increased apoptosis [79]. 4.3.4 Tyrp1

Tyrp1 is the most abundant glycoprotein expressed speciﬁcally in melanocytes. Although the Tyrp1 gene mapping at the brown locus was the ﬁrst member of the tyrosinase gene family to be cloned, its catalytic function (which results in the production of black rather than brown melanin) remains controversial and several lines of evidence suggest that the protein might even serve noncatalytic roles. Despite extensive homology with tyrosinase, Tyrp1 from mouse melanocytic cells binds indolic substrates, but displays an afﬁnity for phenols much lower than tyrosinase. This switch in substrate speciﬁcity could be a consequence of subtle changes within the hydrophobic residues in the respective active sites that have

4.3 Structural and Functional Properties of the Melanogenic Enzymes

been extensively discussed elsewhere [30]. In keeping with these binding properties, the protein puriﬁed from mouse melanocytes behaves as a low-speciﬁcactivity tyrosinase isoenzyme with both tyrosine hydroxylase and dopa oxidase activities [80, 81]. Moreover, mouse Tyrp1 shows DHICA oxidase activity in vitro and mouse melanocytes mutated at the brown locus do not exhibit detectable DHICA oxidase activity, whereas this activity can be readily detected in wild-type melanocytes [18, 19]. However, a role for Tyrp1 as a DHICA oxidase may not be general, as some crucial differences regarding substrate speciﬁcities between the mouse and human melanogenic enzymes have been reported. Indeed, human Tyrp1 does not display DHICA oxidase activity [82], while human tyrosinase does [20]. This suggests that DHICA metabolism could follow alternative enzymatic pathways in mouse and human melanocytes. Apart from its catalytic potential, Tyrp1 participates with tyrosinase in the formation of multimeric complexes [59], which may be important for the regulation of melanogenesis or for the assembly of a melanogenic apparatus. Tyrp1 assists the correct trafﬁcking of tyrosinase to the melanosomes [60] and mediates a signiﬁcant stabilization of the enzyme [58]. It has been proposed that Tyrp1 would be mostly devoid of enzymatic activity and that its strong stabilizing effect on tyrosinase could be the main role for Tyrp1 in melanogenesis. In agreement with this view, mutation of Tyrp1 causes OCA3, which is associated with moderate hypopigmentation of the skin, hair, and eyes. In OCA3, mutated Tyrp1 is retained within the ER and appropriate processing of normal tyrosinase is also aborted, thus accounting for the observed decrease in the production of melanin pigments [60]. 4.3.5 Other Melanosomal Proteins

In addition to the tyrosinase family proteins, genetic evidence shows that other melanosomal proteins are needed for normal melanogenesis, although they may lack a deﬁned enzymatic activity and may serve structural or regulatory roles. The relevance of these proteins for melanogenesis is highlighted by the pigmentation phenotypes of mice mutated at the corresponding loci. We will brieﬂy mention three of them, silver/gp100/gp87/Pmel-17, P (pink-eyed dilution), and MATP (membrane-associated transporter protein)/AIM-1/underwhite/SCL45A2, because these proteins appear to modulate either tyrosinase activity or the rate of distal steps in melanogenesis. The Pmel-17 gene maps at the silver locus whose mutation causes a pigmentation phenotype in mice characterized by premature graying [83]. The silver protein undergoes proteolytic processing within the eumelanosome that provokes its aggregation and contributes to the formation of the lamellar network of the organelle [84], which may trigger the deposit of melanin [85]. Moreover, the possible involvement of the silver protein in facilitating the polymerization of DHICA to melanin has been proposed [22, 23], but so far no precise enzymatic activity has been demonstrated for this protein.

101

102

4 Biosynthesis of Melanins

Mutations of the P and MATP/AIM-1/underwhite/SCL45A2 proteins lead to OCA2 and OCA4 oculocutaneous albinism, respectively. Both proteins display the 12 transmembrane fragments structural arrangement typical of membrane transporters, but their substrate speciﬁcity has not yet been deﬁnitively demonstrated. It has been postulated that the P protein might be an anionic transporter involved in the control of the melanosomal pH, which is critical for tyrosinase activity and processing, and therefore melanogenesis [86]. Regarding MATP, although its substrate for transport is still unknown, in OCA4 melanocytes tyrosinase processing and intracellular trafﬁcking to the melanosome are also disrupted and, consequently, the normal maturation of these organelles is impaired [87].

4.4 Regulation of the Melanogenic Pathway

Mammalian melanogenesis is a highly regulated pathway, as shown by the large number of genes that impinge on the process [88]. Regulatory events may take place in virtually any aspect of melanocyte biology, including melanocyte development and differentiation, the expression of melanosomal components, organelle biogenesis and/or transport, control of pigment-type switching, and others [89]. We discuss here only mechanisms that act directly on the melanogenic pathway to regulate either the type or the amount of pigment formed. 4.4.1 Eumelanogenesis versus Pheomelanogenesis: Regulation of the Type of Melanin Pigments

A wealth of genetic, biochemical and pharmacological evidence has established that the main factor governing the type of pigment formed in melanocytes is signaling from the melanocortin 1 receptor (MC1R). MC1R belongs to a ﬁvemember subgroup of class A G-protein-coupled receptors designated MC1R to MC5R. Their speciﬁc ligands, called melanocortins, are peptide hormones formed by proteolytic cleavage of proopiomelanocortin. MC1R is expressed preferentially but not exclusively in epidermal melanocytes [90, 91] and its speciﬁc ligands are α-melanocyte-stimulating hormone (α-MSH) and adrenocorticotrophin (ACTH). Importantly, these ligands are synthesized not only in the pituitary gland but also by skin keratinocytes, thus allowing for a local paracrine regulation [92] that may be critically involved in physiological adaptations of the skin such as the tanning response [93]. Following activation by α-MSH or ACTH, MC1R stimulates cAMP synthesis. cAMP, the principal intracellular mediator of α-MSH [94], promotes the melanogenic response to the hormones with an increase in tyrosinase activity and a switch from production of light-colored and poorly photoprotective pheomelanins to synthesis of darker and more photoprotective eumelanins [24]. Accordingly, high MC1R signaling in mice melanocytes due to high levels of agonists [95] or gain-of-function mutations [96] leads to eumelanogenesis and dark coats.

4.4 Regulation of the Melanogenic Pathway

Conversely, absent or poor MC1R signaling due to loss-of-function mutations or expression of the endogenous inhibitor agouti signaling protein (ASIP) leads to pheomelanogenesis [97], with the MC1R null phenotype corresponding to a yellow fur color. In humans, a pigment type switch similar to the one in mice is not observed, but there is no doubt that MC1R signaling is also a major determinant of the type of pigment produced. Human MC1R is an extremely polymorphic gene, with more than 100 natural nonsynonymous variants reported to date [98]. Several hypomorphic MC1R alleles are strongly associated with a phenotype known as RHC (for red hair color), consisting of pheomelanin-rich red hair, abundant freckles, inefﬁcient tanning upon sun exposure, high sensitivity to UV radiationinduced skin damage, and increased skin cancer risk. Moreover, a carrier of two MC1R mutant alleles likely corresponding to complete loss-of-function forms of the protein, and hence considered a MC1R null individual, showed a typical RHC phenotype, thus suggesting that red hair and fair skin is the null human MC1R phenotype [99]. Although no chemical characterization of the pigment produced by this likely MC1R-null individual was reported, there is little doubt as to a preferentially pheomelanic pattern. Whereas the positive relationship between MC1R signaling and eumelanogenesis as opposed to pheomelanogenesis is well established, the molecular events responsible for the stimulation of eumelanogenesis following activation of the MC1R by its peptide ligands are only partially understood. Given that pheomelanogenesis depends upon the conjugation of l-dopaquinone and the thiol group of sulfur-containing compounds such as Cys, the simplest possible scenario would posit that under conditions of low tyrosinase activity, the supply of thiol compounds would continuously exceed the low rate of l-dopaquinone formation. Accordingly, essentially all the l-dopaquinone formed within melanosomes would be trapped by conjugation with low molecular weight thiolic compounds, thus directing the melanogenic pathway towards pheomelanogenesis. Conversely, MC1R activation following binding of melanocortin peptides would stimulate tyrosinase. Eventually, a threshold of enzymatic activity would be reached where the rate of production of l-dopaquinone would exceed the supply of thiolic compounds. In these circumstances, this excess of l-dopaquinone would be allowed to evolve to dopachrome and would eventually be incorporated into a growing eumelanin polymer. In support of this simple model, stimulation of mouse melanocytes with α-MSH ﬁrst increases pheomelanin levels, prior to the synthesis of eumelanins, whereas ASIP decreased the production of both types of pigment, probably as a result of downregulation of tyrosinase [100]. Although it is clear that the melanocortin-dependent stimulation of eumelanogenesis in rodent melanocytes and melanoma cells occurs in large part via the increase of tyrosinase activity, the relative contributions of transcriptional and post-transcriptional mechanisms remain uncertain. Transcriptional stimulation is mainly achieved through the cAMP dependent activation of the expression of Microphthalmia (MITF), a master regulator of melanocyte biology that activates several melanogenic genes [101]. MITF binds to speciﬁc target sequences in the

103

104

4 Biosynthesis of Melanins

promoter of the tyrosinase gene, resulting in increased levels of tyrosinase mRNA with a subsequent increase in tyrosinase abundance. However, in human melanocytes stimulated with α-MSH the increases in tyrosinase abundance are modest and it has been shown that intracellular cAMP elevating agents such as forskolin increase MITF protein levels without a concomitant increase in tyrosinase mRNA, thus suggesting the occurrence of post-transcriptional regulatory mechanisms [102]. Moreover, careful studies have established that even in mouse melanocytes the observed increase in tyrosinase mRNA abundance and protein levels is usually lower that the stimulation of tyrosinase speciﬁc activity, thus showing that in addition to transcriptional effects, α-MSH triggers a post-translational stimulation of tyrosinase [103]. The possible molecular basis of these post-translational events will be brieﬂy discussed below. Changes in the levels and/or activity of other proteins in addition to tyrosinase are thought to contribute to the switch of pigment type production in response to α-MSH. Interestingly, in addition to tyrosinase, Dct/Tyrp2 gene expression is upregulated by α-MSH and downregulated by ASIP [104, 105], thus suggesting a role for Dct/Tyrp2 as a positive regulator of eumelanogenesis. In keeping with this possibility, Dct activity and the relative levels of eumelanins and pheomelanins are signiﬁcantly decreased in mouse melanocytes carrying the slaty or slaty light mutations in the Dct locus [106]. Moreover, other factors such as the availability of thiol compounds might be modulated by α-MSH and may contribute to the regulation of the pigment type produced in melanocytes. 4.4.2 Regulation of the Amount of Pigment

Tyrosinase-catalyzed hydroxylation of tyrosine is the rate-limiting step in melanogenesis, and most often there is an excellent positive correlation between changes in the rate of this reaction measured in situ and variations in the melanin content in melanocyte cultures of different racial origin or phenotype. Tyrosinase activity can be modulated through changes in the intracellular concentration of the enzyme or by modiﬁcation of the speciﬁc activity of pre-existing enzyme molecules. Both types of regulation are operative in mammalian melanocytes. 4.4.2.1 Regulation of Tyrosinase Levels Intracellular protein levels reﬂect the balance between the rates of their synthesis and their degradation. As described above, the biosynthesis of tyrosinase and other melanosomal proteins is stimulated by the melanocortin hormones. On the other hand, the rate of tyrosinase degradation is regulated by both intracellular factors and external signaling molecules. Among the best-characterized intracellular factors contributing to the control of tyrosinase degradation, we will brieﬂy discuss mutations in the tyrosinase gene affecting the structure of the protein [60], the intracellular composition of fatty acids [107–109], and interactions with other melanosomal proteins [58, 59]. Concerning external signaling molecules, the tyrosinase-destabilizing effects of cytokines normally present in the skin have been reported [110, 111].

4.4 Regulation of the Melanogenic Pathway

Intracellular trafﬁc of melanosome-resident proteins and other proteins of the biosynthetic-secretory pathway is regulated by stringent quality control mechanisms, whereby misfolded or incorrectly assembled proteins are retained in the ER. Misfolded proteins accumulating in the ER are frequently retrotranslocated to the cytosol and degraded by proteasomes [112]. Proteasomal degradation following retrotranslocation involves ubiquitylation, a post-translational modiﬁcation consisting of the covalent attachment of the 76-amino-acid ubiquitin polypeptide to the ε-amino group of a Lys residue in a protein substrate. A number of diseasecausing mutations in the tyrosinase gene found in OCA1 patients result in extensive intracellular retention in the ER and subsequent proteasomal degradation of tyrosinase [60]. This process involves aberrant N-glycosylation of the enzyme causing abnormal interaction with ER-resident chaperones [47, 113–115]. Aberrant processing with ubiquitylation and degradation of tyrosinase is also causally related to the depigmented phenotype of human melanomas [51] and also results in lack of pigmentation in normal melanocytes [56], thus showing that increased tyrosinase degradation can have profound effects on visible pigmentation. Recent ﬁndings suggest that the ubiquitin–proteasome system (UPS) plays a much more complex and sophisticated role in tyrosinase regulation than merely preventing the accumulation of misfolded and enzymatically inactive protein molecules in carriers of OCA1 mutations or in amelanotic melanomas (reviewed in [116]). Indeed, proteasomal degradation of wild-type tyrosinase appears also possible and susceptible of regulation. Fatty acids have been shown to modulate the stability of tyrosinase by modifying the activity of the UPS in opposing directions, depending on their structure. Thus, saturated palmitic acid stabilizes tyrosinase, whereas the monounsaturated and longer linoleic acid increases tyrosinase ubiquitylation and accelerates its degradation [108]. Accordingly, linoleic acid has been shown to exert a pharmacologic hypopigmenting effect [117]. More recently, it has been reported that the rate of tyrosinase ubiquitylation and hence proteasomal degradation of the enzyme is controlled at least partially by the activity of p38 mitogen-activated protein kinase (MAPK). p38 MAPK is activated by a variety of internal or environmental stresses including UV radiation and might in fact play a role in tyrosinase transcriptional activation during the tanning response [118]. Intriguingly, inhibition of p38 MAPK expression decreased tyrosinase ubiquitylation and consequently stimulated melanogenesis, suggesting that p38 might in fact behave as a negative regulator of tyrosinase intracellular stability [119]. p38 MAPK is activated by α-MSH in mouse melanocytes [120] and in human melanoma cells (our unpublished results), and is also activated following exposure to UV radiation [118]. This suggests that regulated tyrosinase ubiquitylation and proteasomal degradation can provide a mechanism to ﬁne-tune the stimulatory effect of melanogenic stimuli such as the melanocortin peptides or UV radiation, whereby an excessive melanogenic response would be prevented via accelerated degradation of tyrosinase. It has been reported that tyrosinase quality control and degradation can also occur in a post-Golgi compartment probably related with the endosomal–lysosomal system, following complete maturation and independently of the UPS machinery

105

106

4 Biosynthesis of Melanins

[121]. This nonproteasomal degradative pathway could be induced by active-site directed inhibitors of tyrosinase such as phenylthiourea and might involve a speciﬁc Cys protease. The relative contributions of the proteasomal and nonproteasomal pathways to the degradation of normal and mutant tyrosinase variants remain to be determined. Using puriﬁed preparations of tyrosinase and Tyrp1 we showed that both proteins interact strongly and speciﬁcally in detergent solution to form heterodimeric complexes [59]. The functional consequences of this speciﬁc interaction have been analyzed. Hearing et al. ﬁrst demonstrated that tyrosinase is less stable in mutant Tyrp1 mouse melanocytes than in cells expressing wild-type Tyrp1 and that forced expression of wild-type Tyrp1 in mutant melanocytes partly rescued the normal phenotype by increasing tyrosinase stability [58]. Direct interaction of tyrosinase and Tyrp1 in vivo, with signiﬁcant stabilization of the former, was further conﬁrmed by chemical cross-linking experiments [122]. The strong effect of Tyrp1 on tyrosinase maturation and stability might provide a basis for the albino phenotype of OCA3 patients, whereby a mutation in Tyrp1 might impair the maturation and stability of tyrosinase thus decreasing tyrosinase activity [60]. In summary, tyrosinase degradation appears to follow at least two different pathways. On the one hand, a UPS-dependent pathway might be particularly relevant for mutant or otherwise misfolded tyrosinase molecules. On the other hand, a post-Golgi proteasome-independent route most likely related with endosomal– lysosomal compartments and involving a speciﬁc Cys protease might act on fully glycosylated mature forms of the protein, and hence might be particularly relevant for the turnover of normal and correctly processed enzyme. The proteasomedependent pathway appears susceptible of regulation by internal factors (fatty acids) as well as by external stimuli (α-MSH, UV radiation) acting at least partially through p38 MAPK. Regulation of the UPS might contribute to the ﬁne-tuning of pigmentation by preventing excessive melanin synthesis in cells following high intensity melanogenic stimuli. Moreover, the skin whitening effects of unsaturated fatty acids in vivo provide a proof of principle showing that pharmacological modulation of tyrosinase stability can be used as a strategy to treat pigmentary disorders. 4.4.2.2 Control of Tyrosinase-Speciﬁc Activity Several possible post-transcriptional mechanisms of regulation of tyrosinase speciﬁc activity have been reported, including activation by protein kinase C-dependent phosphorylation of residues located in the cytosolic tail of the protein [123], and changes in the melanosomal pH. Modulation of the melanosomal pH appears a particularly important mechanism, as it is likely one of the factors underlying racial differences in skin pigmentation and may also contribute to tyrosinase activation following stimulation of melanocytes with α-MSH. Melanocytes of black and Caucasian skin show very similar levels of tyrosinase mRNA and protein abundance, but “in situ” tyrosinase activity measured in intact cells is much higher in melanocytes from black donors [124]. Thus, the good correlation between melanin content and tyrosinase activity in melanocytes of

4.5 Conclusions and Perspectives

different racial origin is not accounted for by parallel differences in enzyme abundance, indicative of a major role for post-transcriptional regulatory mechanisms. This important difference in tyrosinase speciﬁc activity in melanosomes of different racial origin appears related with the pH of the organelle. Melanosomes are organelles of the endosomal–lysosomal lineage and as such they are expected to be acidic. This has been conﬁrmed by estimations of the melanosomal pH in mouse melanocytes and melanoma cells [125, 126], as well as in normal human melanocyte cultures derived from Caucasian donors [127]. Interestingly, the melanosomes of melanocytes from black donors are more neutral than those derived from Caucasian skin. Moreover, treatment of melanocytes with lysosomotropic agents that increase the melanosomal pH such as ammonium chloride or the ionophores nigericin and monensin strongly stimulates tyrosinase activity in melanocytes from Caucasian donors, but has no effect in black melanocytes [127]. Similar increases in tyrosinase activity were obtained with selective inhibitors of vacuolar proton pump V-ATPase [128]. The optimal pH of mammalian tyrosinase is near neutral and tyrosinase-speciﬁc activity decreases dramatically at acidic pH [129, 130]. Accordingly, a model for racial pigmentation based on differences in melanosomal pH has been proposed, whereby the acidic environment in Caucasian melanosomes would keep tyrosinase largely inactive whereas the more neutral pH in black melanosomes would allow for full tyrosinase activity [127]. However, the interpretation of the activatory effect of agents expected to increase the melanosomal pH is complicated by the observation that tyrosinase processing and transport through the ER to the Golgi, and then to melanosomes via the endosomal sorting system, is enhanced in the presence of protonophore or proton pump inhibitors that increase the pH of intracellular organelles [131]. It has been shown that cAMP increases the pH of melanosomes and regulates the expression of several vacuolar ATPases and ion transporters [132]. Therefore, changes in the pH of melanosomes may partially underlie the melanogenic response to the melanocortin peptides. Accordingly, the melanosomal pH would be controlled by genetic determinants responsible for racial skin pigmentation traits, as well as by external signaling molecules such as the melanocortins. In turn, this modulation of the melanosomal pH would determine changes in tyrosinase speciﬁc activity thus contributing to the regulation of the rate of melanin biosynthesis.

4.5 Conclusions and Perspectives

The chemical analysis of natural melanin pigments and the kinetic and molecular characterization of new melanogenic enzymes have led to a profound modiﬁcation of the melanogenic pathway as originally delineated by Raper and Mason. Melanins were originally considered relatively simple polymers containing essentially one type of decarboxylated DHI-derived monomeric units synthesized from

107

108

4 Biosynthesis of Melanins

l-tyrosine by the action of a single enzyme. Now, eumelanins are seen as complex and heterogeneous structures with both decarboxylated and carboxylated (DHICAderived) units formed by the concerted action of three enzymatic proteins, tyrosinase and the closely related Tyrp1 and Dct/Tyrp2. The relative proportions of both types of monomeric units determine the exact physicochemical properties of the pigment and may be highly variable depending on factors such as the activity of the melanogenic enzymes, the availability of substrates, and chemical properties of the melanosomal milieu like its metal ion content and its pH. Accordingly, a wide and quasicontinuous spectrum of physicochemical properties is expected for the natural pigments – a view much more attractive and consistent with our daily perceptions. However, whereas the biochemistry of eumelanogenesis has been basically worked out, current knowledge of the pheomelanogenic pathway remains incomplete. The participation of low-molecular-weight thiol compounds and the major role for 5-cysteynyldopa and alanyl-hydroxy-benzothiazine monomers has been established, but the structure of other biosynthetic intermediates is still unsolved. The involvement of enzymatic activities different from tyrosinase or nonenzymatic proteins such as membrane transporters also remains an open question. Therefore, it can be anticipated that the chemistry of pheomelanogenesis and the biochemical factors governing the type of pigment formed will be major areas of fruitful research in the near future. Moreover, recent evidence suggests that Dct/Tryp2 and maybe also Tyrp1 play intriguing and important roles in the regulation of cell proliferation and survival under certain stress conditions, in addition to their enzymatic activity in the melanogenic pathway. New and exciting ﬁndings concerning these nonmelanogenic roles of the tyrosinase family proteins can also be anticipated. The study of the regulation of mammalian melanogenesis has revealed a complex and sophisticated network of interconnected and cross-talking signals, including: (i) intramelanocytic factors such as fatty acids or the pH of intracellular organelles, (ii) paracrine effectors released by cells in the epidermal unit, (iii) systemic endocrine signals, and (iv) environmental inﬂuences such as UV radiation. All these factors trigger transcriptional and post-transcriptional regulatory mechanisms whose molecular details and relative importance in different physio(patho)logical situations will be worked out. This increased knowledge of the regulation of normal and aberrant melanogenesis will no doubt lead to better therapeutic approaches to the treatment of pigmentary disorders. Several of the signals that regulate melanin biosynthesis, such as UV radiation or the melanocortin hormones, act coordinately on various aspects of the cellular physiology, including DNA metabolism and cell cycle control, organelle biogenesis, subcellular trafﬁcking, and the rate of a highly specialized metabolic pathway. Moreover, this simultaneous and integrated control of different cellular functions within melanocytes leads to a clear, visible and easily quantiﬁable phenotypic output. Accordingly, the biosynthesis of melanins provides a unique model for the holistic analysis of the regulation of metabolic processes.

References

Acknowledgments

Work in the authors’ laboratory is supported by grants SAF2009-10942 from the Comisión Interministerial de Ciencia y Tecnología (CICYT), Spain, FEDER funds (European Community), and 464/2008 from the CARM, Plan de Ciencia y Tecnología 2007/10. We thank Celia Jiménez-Cervantes for continuous help, support, and collaboration. We apologize to the many colleagues whose relevant work has not been explicitly cited due to space limitations.

References 1 Hearing, V.J. and Tsukamoto, K. (1991)

2

3

4

5

6 7

8

9

10

Enzymatic control of pigmentation in mammals. FASEB J., 5, 2902–2909. Claus, H. and Decker, H. (2006) Bacterial tyrosinases. Syst. Appl. Microbiol., 29, 3–14. Liu, G.Y. and Nizet, V. (2009) Color me bad: microbial pigments as virulence factors. Trends Microbiol., 17, 406–413. Mayer, A.M. (2006) Polyphenol oxidases in plants and fungi: going places? A review. Phytochemistry, 67, 2318–2331. Halaouli, S., Asther, M., Sigoillot, J.C., Hamdi, M., and Lomascolo, A. (2006) Fungal tyrosinases: new prospects in molecular characteristics, bioengineering and biotechnological applications. J. Appl. Microbiol., 100, 219–232. Raper, H.S. (1928) The aerobic oxidases. Physiol. Rev., 8, 245–258. Mason, H.S. (1948) The chemistry of melanin III. Mechanism of the oxidation of dihydroxyphenylalanine by tyrosinase. J. Biol. Chem., 172, 83–90. Kwon, B.S. (1993) Pigmentation genes: the tyrosinase gene family and the pmel 17 gene family. J. Invest. Dermatol., 100, 134S–140S. Nordlund, J.J., Boissy, R.E., Hearing, V.J., King, R.A., Oetting, W.S., and Ortonne, J.P. (eds) (2006) The Pigmentary System: Physiology and Pathophysiology, 2nd edn, Blackwell, Malden, MA. Wilcox, D.E., Porras, A.G., Hwang, Y.T., Lerch, K., Winkler, M.E., and Solomon, E.I. (1985) Substrate analog binding to the coupled binuclear copper active site in tyrosinase. J. Am. Chem. Soc., 107, 4015–4027.

11 García-Canovas, F., Garcia-Carmona, F.,

12

13

14 15

16

17

Sanchez, J.V., Iborra, J.L., and Lozano, J.A. (1982) The role of pH in the melanin biosynthesis pathway. J. Biol. Chem., 257, 8738–8744. Lerner, A.B. and Fitzpatrick, T.B. (1950) Biochemistry of melanin formation. Physiol. Rev., 30, 91–126. Rodríguez-Lopez, J.N., Banon-Arnao, M., Martinez-Ortiz, F., Tudela, J., Acosta, M., Varon, R., and GarciaCanovas, F. (1992) Catalytic oxidation of 2,4,5-trihydroxyphenylalanine by tyrosinase: identiﬁcation and evolution of intermediates. Biochim. Biophys. Acta, 1160, 221–228. Pawelek, J.M. (1991) After dopachrome? Pigment Cell Res., 4, 53–62. Aroca, P., Garcia-Borron, J.C., Solano, F., and Lozano, J.A. (1990) Regulation of mammalian melanogenesis. I: partial puriﬁcation and characterization of a dopachrome converting factor: dopachrome tautomerase. Biochim. Biophys. Acta, 1035, 266–275. Palumbo, A., Solano, F., Misuraca, G., Aroca, P., Garcia-Borron, J.C., Lozano, J.A., and Prota, G. (1991) Comparative action of dopachrome tautomerase and metal ions on the rearrangement of dopachrome. Biochim. Biophys. Acta, 1115, 1–5. Aroca, P., Solano, F., Salinas, C., Garcia-Borron, J.C., and Lozano, J.A. (1992) Regulation of the ﬁnal phase of mammalian melanogenesis. The role of dopachrome tautomerase and the ratio between 5,6-dihydroxyindole-2carboxylic acid and 5,6-dihydroxyindole. Eur. J. Biochem., 208, 155–163.

109

110

4 Biosynthesis of Melanins 18 Jimenez-Cervantes, C., Solano, F.,

19

20

21

22

23

24

25

Kobayashi, T., Urabe, K., Hearing, V.J., Lozano, J.A., and Garcia-Borron, J.C. (1994) A new enzymatic function in the melanogenic pathway. The 5,6-dihydroxyindole-2-carboxylic acid oxidase activity of tyrosinase-related protein-1 (TRP1). J. Biol. Chem., 269, 17993–18001. Kobayashi, T., Urabe, K., Winder, A., Jimenez-Cervantes, C., Imokawa, G., Brewington, T., Solano, F., GarciaBorron, J.C., and Hearing, V.J. (1994) Tyrosinase related protein 1 (TRP1) functions as a DHICA oxidase in melanin biosynthesis. EMBO J., 13, 5818–5825. Olivares, C., Jimenez-Cervantes, C., Lozano, J.A., Solano, F., and GarciaBorron, J.C. (2001) The 5,6-dihydroxyindole-2-carboxylic acid (DHICA) oxidase activity of human tyrosinase. Biochem J., 354, 131–139. Urabe, K., Aroca, P., Tsukamoto, K., Mascagna, D., Palumbo, A., Prota, G., and Hearing, V.J. (1994) The inherent cytotoxicity of melanin precursors: a revision. Biochim. Biophys. Acta, 1221, 272–278. Chakraborty, A.K., Platt, J.T., Kim, K.K., Kwon, B.S., Bennett, D.C., and Pawelek, J.M. (1996) Polymerization of 5,6-dihydroxyindole-2-carboxylic acid to melanin by the pmel 17/silver locus protein. Eur. J. Biochem., 236, 180–188. Lee, Z.H., Hou, L., Moellmann, G., Kuklinska, E., Antol, K., Fraser, M., Halaban, R., and Kwon, B.S. (1996) Characterization and subcellular localization of human Pmel 17/silver, a 110-kDa (pre)melanosomal membrane protein associated with 5,6-dihydroxyindole-2-carboxylic acid (DHICA) converting activity. J. Invest. Dermatol., 106, 605–610. Ito, S. and Wakamatsu, K. (2003) Quantitative analysis of eumelanin and pheomelanin in humans, mice, and other animals: a comparative review. Pigment Cell Res., 16, 523–531. Jara, J.R., Aroca, P., Solano, F., Martinez, J.H., and Lozano, J.A. (1988) The role of sulfhydryl compounds in mammalian melanogenesis: the effect of

26

27

28

29

30

31

32

33

34

35

cysteine and glutathione upon tyrosinase and the intermediates of the pathway. Biochim. Biophys. Acta, 967, 296–303. Prota, G. (1988) Progress in the chemistry of melanins and related metabolites. Med. Res. Rev., 8, 525–556. Prota, G. (1992) Pheomelanins and trichochromes, in Melanins and Melanogenesis, Academic Press, San Diego, CA, pp. 134–152. Agrup, G., Falck, B., Kennedy, B.M., Rorsman, H., Rosengren, A.M., and Rosengren, E. (1975) Formation of cysteinyldopa from glutathionedopa in melanoma. Acta Derm. Venereol., 55, 1–3. Jackson, I.J., Budd, P., Horn, J.M., Johnson, R., Raymond, S., and Steel, K. (1994) Genetics and molecular biology of mouse pigmentation. Pigment Cell Res., 7, 73–80. García-Borrón, J.C. and Solano, F. (2002) Molecular anatomy of tyrosinase and its related proteins: beyond the histidine-bound metal catalytic center. Pigment Cell Res., 15, 162–173. Lerner, A.B., Fitzpatrick, T.B., Calkins, E., and Summerson, W.H. (1950) Mammalian tyrosinase. The relationship of copper to enzymatic activity. J. Biol. Chem., 187, 793–802. Solano, F., Jimenez-Cervantes, C., Martinez-Liarte, J.H., Garcia-Borron, J.C., Jara, J.R., and Lozano, J.A. (1996) Molecular mechanism for catalysis by a new zinc-enzyme, dopachrome tautomerase. Biochem. J., 313, 447–453. Furumura, M., Solano, F., Matsunaga, N., Sakai, C., Spritz, R.A., and Hearing, V.J. (1998) Metal ligand-binding speciﬁcities of the tyrosinase-related proteins. Biochem. Biophys. Res. Commun., 242, 579–585. Oetting, W.S. and King, R.A. (1994) Analysis of tyrosinase mutations associated with tyrosinase-related oculocutaneous albinism (OCA1). Pigment Cell Res., 7, 285–290. Olivares, C., Garcia-Borron, J.C., and Solano, F. (2002) Identiﬁcation of active site residues involved in metal cofactor binding and stereospeciﬁc substrate recognition in mammalian tyrosinase.

References

36

37

38

39

40

41

42

43

44

45

Implications to the catalytic cycle. Biochemistry, 41, 679–686. Klabunde, T., Eicken, C., Sacchettini, J.C., and Krebs, B. (1998) Crystal structure of a plant catechol oxidase containing a dicopper center. Nat. Struct. Biol., 5, 1084–1090. Matoba, Y., Kumagai, T., Yamamoto, A., Yoshitsu, H., and Sugiyama, M. (2006) Crystallographic evidence that the dinuclear copper center of tyrosinase is ﬂexible during catalysis. J. Biol. Chem., 281, 8981–8990. Solomon, E.I. and Lowery, M.D. (1993) Electronic structure contributions to function in bioinorganic chemistry. Science, 259, 1575–1581. Lerch, K. (1983) Neurospora tyrosinase: structural, spectroscopic and catalytic properties. Mol. Cell. Biochem., 52, 125–138. Nakamura, M., Nakajima, T., Ohba, Y., Yamauchi, S., Lee, B.R., and Ichishima, E. (2000) Identiﬁcation of copper ligands in Aspergillus oryzae tyrosinase by site-directed mutagenesis. Biochem. J., 350, 537–545. Hearing, V.J. and Jimenez, M. (1989) Analysis of mammalian pigmentation at the molecular level. Pigment Cell Res., 2, 75–85. Spritz, R.A., Ho, L., Furumura, M., and Hearing, V.J.J. (1997) Mutational analysis of copper binding by human tyrosinase. J. Invest. Dermatol., 109, 207–212. Jimbow, K., Hua, C., Gomez, P.F., Hirosaki, K., Shinoda, K., Salopek, T.G., Matsusaka, H., Jin, H.Y., and Yamashita, T. (2000) Intracellular vesicular trafﬁcking of tyrosinase gene family protein in eu- and pheomelanosome biogenesis. Pigment Cell Res., 13 (Suppl. 8), 110–117. Jimbow, K., Park, J.S., Kato, F., Hirosaki, K., Toyofuku, K., Hua, C., and Yamashita, T. (2000) Assembly, target-signaling and intracellular transport of tyrosinase gene family proteins in the initial stage of melanosome biogenesis. Pigment Cell Res., 13, 222–229. Setaluri, V. (2000) Sorting and targeting of melanosomal membrane proteins:

46

47

48

49

50

51

52

53

54

signals, pathways, and mechanisms. Pigment Cell Res., 13, 128–134. Jackson, I.J. (1994) Molecular and developmental genetics of mouse coat color. Annu. Rev. Genet., 28, 189–217. Branza-Nichita, N., Negroiu, G., Petrescu, A.J., Garman, E.F., Platt, F.M., Wormald, M.R., Dwek, R.A., and Petrescu, S.M. (2000) Mutations at critical N-glycosylation sites reduce tyrosinase activity by altering folding and quality control. J. Biol. Chem., 275, 8169–8175. Harrison, M.D., Jones, C.E., Solioz, M., and Dameron, C.T. (2000) Intracellular copper routing: the role of copper chaperones. Trends Biochem. Sci., 25, 29–32. Olivares, C., Solano, F., and GarciaBorron, J.C. (2003) Conformationdependent post-translational glycosylation of tyrosinase. Requirement of a speciﬁc interaction involving the CuB metal binding site. J. Biol. Chem., 278, 15735–15743. Hearing, V.J., Ekel, T.M., and Montague, P.M. (1981) Mammalian tyrosinase: isozymic forms of the enzyme. Int. J. Biochem., 13, 99–103. Halaban, R., Cheng, E., Zhang, Y., Moellmann, G., Hanlon, D., Michalak, M., Setaluri, V., and Hebert, D.N. (1997) Aberrant retention of tyrosinase in the endoplasmic reticulum mediates accelerated degradation of the enzyme and contributes to the dedifferentiated phenotype of amelanotic melanoma cells. Proc. Natl. Acad. Sci. USA, 94, 6210–6215. Raposo, G. and Marks, M.S. (2007) Melanosomes – dark organelles enlighten endosomal membrane transport. Nat. Rev. Mol. Cell Biol., 8, 786–797. Setty, S.R., Tenza, D., Sviderskaya, E.V., Bennett, D.C., Raposo, G., and Marks, M.S. (2008) Cell-speciﬁc ATP7A transport sustains copper-dependent tyrosinase activity in melanosomes. Nature, 454, 1142–1146. Petris, M.J., Strausak, D., and Mercer, J.F. (2000) The Menkes copper transporter is required for the activation

111

112

4 Biosynthesis of Melanins

55

56

57

58

59

60

61

62

63

of tyrosinase. Hum. Mol. Genet., 9, 2845–2851. Oetting, W.S. (2000) The tyrosinase gene and oculocutaneous albinism type 1 (OCA1): a model for understanding the molecular biology of melanin formation. Pigment Cell Res., 13, 320–325. Toyofuku, K., Wada, I., Spritz, R.A., and Hearing, V.J. (2001) The molecular basis of oculocutaneous albinism type 1 (OCA1): sorting failure and degradation of mutant tyrosinases results in a lack of pigmentation. Biochem. J., 355, 259–269. Oetting, W.S., Fryer, J.P., Shriram, S., and King, R.A. (2003) Oculocutaneous albinism type 1: the last 100 years. Pigment Cell Res., 16, 307–311. Kobayashi, T., Imokawa, G., Bennett, D.C., and Hearing, V.J. (1998) Tyrosinase stabilization by Tyrp1 (the brown locus protein). J. Biol. Chem., 273, 31801–31805. Jimenez-Cervantes, C., MartinezEsparza, M., Solano, F., Lozano, J.A., and Garcia-Borron, J.C. (1998) Molecular interactions within the melanogenic complex: formation of heterodimers of tyrosinase and TRP1 from B16 mouse melanoma. Biochem. Biophys. Res. Commun., 253, 761–767. Toyofuku, K., Wada, I., Valencia, J.C., Kushimoto, T.O., Ferrana, V.J., and Hearing, V.J. (2001) Oculocutaneous albinism types 1 and 3 are ER retention diseases: mutation of tyrosinase or Tyrp1 can affect the processing of both mutant and wild-type proteins. FASEB J., 15, 2149–2161. Schweikardt, T., Olivares, C., Solano, F., Jaenicke, E., Garcia-Borron, J.C., and Decker, H. (2007) A three-dimensional model of mammalian tyrosinase active site accounting for loss of function mutations. Pigment Cell Res., 20, 394–401. Olivares, C. and Solano, F. (2009) New insights into the active site structure and catalytic mechanism of tyrosinase and its related proteins. Pigment Cell Melanoma Res, 22, 750–760. Pomerantz, S.H. (1966) The tyrosine hydroxylase activity of mammalian tyrosinase. J. Biol. Chem., 241, 161–168.

64 Pomerantz, S.H. and Warner, M.C.

65

66

67

68

69

70

71

72

73

(1967) 3,4-dihydroxy-l-phenylalanine as the tyrosinase cofactor. Occurrence in melanoma and binding constant. J. Biol. Chem., 242, 5308–5314. Inoue, T., Shiota, Y., and Yoshizawa, K. (2008) Quantum chemical approach to the mechanism for the biological conversion of tyrosine to dopaquinone. J. Am. Chem. Soc., 130, 16890–16897. Cooksey, C.J., Garratt, P.J., Land, E.J., Pavel, S., Ramsden, C.A., Riley, P.A., and Smit, N.P. (1997) Evidence of the indirect formation of the catecholic intermediate substrate responsible for the autoactivation kinetics of tyrosinase. J. Biol. Chem., 272, 26226–26235. Land, E.J., Ramsden, C.A., and Riley, P.A. (2003) Tyrosinase autoactivation and the chemistry of ortho-quinone amines. Acc. Chem. Res., 36, 300–308. Jara, J.R., Solano, F., and Lozano, J.A. (1988) Assays for mammalian tyrosinase: a comparative study. Pigment Cell Res., 1, 332–339. Land, E.J., Ramsden, C.A., and Riley, P.A. (2007) The mechanism of suicide-inactivation of tyrosinase: a substrate structure investigation. Tohoku J. Exp. Med., 212, 341–348. Land, E.J., Ramsden, C.A., Riley, P.A., and Stratford, M.R. (2008) Evidence consistent with the requirement of cresolase activity for suicide inactivation of tyrosinase. Tohoku J. Exp. Med., 216, 231–238. Jackson, I.J., Chambers, D.M., Tsukamoto, K., Copeland, N.G., Gilbert, D.J., Jenkins, N.A., and Hearing, V. (1992) A second tyrosinase-related protein, TRP-2, maps to and is mutated at the mouse slaty locus. EMBO J., 11, 527–535. Raposo, G., Tenza, D., Murphy, D.M., Berson, J.F., and Marks, M.S. (2001) Distinct protein sorting and localization to premelanosomes, melanosomes and lysosomes in pigmented melanocytic cells. J. Cell. Biol., 152, 809–824. Aroca, P., Martinez-Liarte, J.H., Solano, F., Garcia-Borron, J.C., and Lozano, J.A. (1992) The action of glycosylases on dopachrome (2-carboxy-2,3-

References

74

75

76

77

78

79

80

81

82

dihydroindole-5,6-quinone) tautomerase. Biochem. J., 284, 109–113. Solano, F., Martinez-Liarte, J.H., Jimenez-Cervantes, C., Garcia-Borron, J.C., and Lozano, J.A. (1994) Dopachrome tautomerase is a zinccontaining enzyme. Biochem. Biophys. Res. Commun., 204, 1243–1250. Odh, G., Hindemith, A., Rosengren, A.M., Rosengren, E., and Rorsman, H. (1993) Isolation of a new tautomerase monitored by the conversion of d-dopachrome to 5,6-dihydroxyindole. Biochem. Biophys. Res. Commun., 197, 619–624. Palumbo, A., d’Ischia, M., Misuraca, G., De Martino, L., and Prota, G. (1994) A new dopachrome-rearranging enzyme from the ejected ink of the cuttleﬁsh Sepia ofﬁcinalis. Biochem. J., 299, 839–844. Aroca, P., Solano, F., Garcia-Borron, J.C., and Lozano, J.A. (1991) Speciﬁcity of dopachrome tautomerase and inhibition by carboxylated indoles. Considerations on the enzyme active site. Biochem. J., 277, 393–397. Steel, K.P., Davidson, D.R., and Jackson, I.J. (1992) TRP-2/DT, a new early melanoblast marker, shows that steel growth factor (c-kit ligand) is a survival factor. Development, 115, 1111–1119. Sendoel, A., Kohler, I., Fellmann, C., Lowe, S.W., and Hengartner, M.O. (2010) HIF-1 antagonizes p53-mediated apoptosis through a secreted neuronal tyrosinase. Nature, 465, 577–583. Jimenez-Cervantes, C., Garcia-Borron, J.C., Valverde, P., Solano, F., and Lozano, J.A. (1993) Tyrosinase isoenzymes in mammalian melanocytes. 1. Biochemical characterization of two melanosomal tyrosinases from B16 mouse melanoma. Eur. J. Biochem., 217, 549–556. Jimenez, M., Tsukamoto, K., and Hearing, V.J. (1991) Tyrosinases from two different loci are expressed by normal and by transformed melanocytes. J. Biol. Chem., 266, 1147–1156. Boissy, R.E., Sakai, C., Zhao, H., Kobayashi, T., and Hearing, V.J. (1998)

83

84

85

86

87

88

89

90

91

Human tyrosinase related protein-1 (TRP-1) does not function as a DHICA oxidase activity in contrast to murine TRP-1. Exp. Dermatol., 7, 198–204. Martinez-Esparza, M., JimenezCervantes, C., Bennett, D.C., Lozano, J.A., Solano, F., and Garcia-Borron, J.C. (1999) The mouse silver locus encodes a single transcript truncated by the silver mutation. Mamm. Genome, 10, 1168–1171. Kushimoto, T., Basrur, V., Valencia, J., Matsunaga, J., Vieira, W.D., Ferrans, V.J., Muller, J., Appella, E., and Hearing, V.J. (2001) A model for melanosome biogenesis based on the puriﬁcation and analysis of early melanosomes. Proc. Natl. Acad. Sci. USA, 98, 10698–10703. Solano, F., Martinez-Esparza, M., Jimenez-Cervantes, C., Hill, S.P., Lozano, J.A., and Garcia-Borron, J.C. (2000) New insights on the structure of the mouse silver locus and on the function of the silver protein. Pigment Cell Res., 13 (Suppl. 8), 118–124. Chen, K., Manga, P., and Orlow, S.J. (2002) Pink-eyed dilution protein controls the processing of tyrosinase. Mol. Biol. Cell., 13, 1953–1964. Costin, G.E., Valencia, J.C., Vieira, W.D., Lamoreux, M.L., and Hearing, V.J. (2003) Tyrosinase processing and intracellular trafﬁcking is disrupted in mouse primary melanocytes carrying the underwhite (uw) mutation. A model for oculocutaneous albinism (OCA) type 4. J. Cell. Sci., 116, 3203–3212. Bennett, D.C. and Lamoreux, M.L. (2003) The color loci of mice – a genetic century. Pigment Cell Res., 16, 333–344. Lamoreux, M.L., Delmas, V., Larue, L., and Bennett, D.C. (2010) The Colors of Mice: A Model Genetic Network, Wiley-Blackwell, Oxford. Roberts, D.W., Newton, R.A., Beaumont, K.A., Helen, L.J., and Sturm, R.A. (2006) Quantitative analysis of MC1R gene expression in human skin cell cultures. Pigment Cell Res., 19, 76–89. Bohm, M., Luger, T.A., Tobin, D.J., and Garcia-Borron, J.C. (2006) Melanocortin receptor ligands: new horizons for skin

113

114

4 Biosynthesis of Melanins

92

93

94

95

96

97

98

99

100

biology and clinical dermatology. J. Invest. Dermatol., 126, 1966–1975. Slominski, A., Tobin, D.J., Shibahara, S., and Wortsman, J. (2004) Melanin pigmentation in mammalian skin and its hormonal regulation. Physiol. Rev., 84, 1155–1228. Cui, R., Widlund, H.R., Feige, E., Lin, J.Y., Wilensky, D.L., Igras, V.E., D’Orazio, J., Fung, C.Y., Schanbacher, C.F., Granter, S.R., and Fisher, D.E. (2007) Central role of p53 in the suntan response and pathologic hyperpigmentation. Cell, 128, 853–864. Busca, R. and Ballotti, R. (2000) Cyclic AMP a key messenger in the regulation of skin pigmentation. Pigment Cell Res., 13, 60–69. Geschwind, I.I. (1966) Change in hair color in mice induced by injection of alpha-MSH. Endocrinology, 79, 1165–1167. Robbins, L.S., Nadeau, J.H., Johnson, K.R., Kelly, M.A., Roselli-Rehfuss, L., Baack, E., Mountjoy, K.G., and Cone, R.D. (1993) Pigmentation phenotypes of variant extension locus alleles result from point mutations that alter MSH receptor function. Cell, 72, 827–834. Bultman, S.J., Michaud, E.J., and Woychik, R.P. (1992) Molecular characterization of the mouse agouti locus. Cell, 71, 1195–1204. Perez Oliva, A.B., Fernendez, L.P., Detorre, C., Herraiz, C., MartinezEscribano, J.A., Benitez, J., Lozano Teruel, J.A., Garcia-Borron, J.C., Jimenez-Cervantes, C., and Ribas, G. (2009) Identiﬁcation and functional analysis of novel variants of the human melanocortin 1 receptor found in melanoma patients. Hum. Mutat., 30, 811–822. Beaumont, K.A., Shekar, S.N., Cook, A.L., Duffy, D.L., and Sturm, R.A. (2008) Red hair is the null phenotype of MC1R. Hum. Mutat., 29, E88–E94. Le Pape, E., Wakamatsu, K., Ito, S., Wolber, R., and Hearing, V.J. (2008) Regulation of eumelanin/pheomelanin synthesis and visible pigmentation in melanocytes by ligands of the melanocortin 1 receptor. Pigment Cell Melanoma Res., 21, 477–486.

101 Levy, C., Khaled, M., and Fisher, D.E.

102

103

104

105

106

107

108

(2006) MITF: master regulator of melanocyte development and melanoma oncogene. Trends Mol. Med., 12, 406–414. Newton, R.A., Cook, A.L., Roberts, D.W., Leonard, J.H., and Sturm, R.A. (2007) Post-transcriptional regulation of melanin biosynthetic enzymes by cAMP and resveratrol in human melanocytes. J. Invest. Dermatol., 127, 2216–2227. Fuller, B.B., Lunsford, J.B., and Iman, D.S. (1987) Alpha-melanocytestimulating hormone regulation of tyrosinase in Cloudman S-91 mouse melanoma cell cultures. J. Biol. Chem., 262, 4024–4033. 104. Rouzaud, F. and Hearing, V.J. (2006) Analysis of the transcriptional regulation of melanocytic genes by alphaMSH using the cDNA microarray technique. Cell. Mol. Biol., 52, 21–31. Furumura, M., Sakai, C., Potterf, S.B., Vieira, W.D., Barsh, G.S., and Hearing, V.J. (1998) Characterization of genes modulated during pheomelanogenesis using differential display. Proc. Natl. Acad. Sci. USA, 95, 7374–7378. Costin, G.E., Valencia, J.C., Wakamatsu, K., Ito, S., Solano, F., Milac, A.L., Vieira, W.D., Yamaguchi, Y., Rouzaud, F., Petrescu, A.J., Lamoreux, M.L., and Hearing, V.J. (2005) Mutations in dopachrome tautomerase (Dct) affect eumelanin/pheomelanin synthesis, but do not affect intracellular trafﬁcking of the mutant protein. Biochem. J., 391, 249–259. Ando, H., Funasaka, Y., Oka, M., Ohashi, A., Furumura, M., Matsunaga, J., Matsunaga, N., Hearing, V.J., and Ichihashi, M. (1999) Possible involvement of proteolytic degradation of tyrosinase in the regulatory effect of fatty acids on melanogenesis. J. Lipid Res., 40, 1312–1316. Ando, H., Watabe, H., Valencia, J.C., Yasumoto, K., Furumura, M., Funasaka, Y., Oka, M., Ichihashi, M., and Hearing, V.J. (2004) Fatty acids regulate pigmentation via proteasomal degradation of tyrosinase: a new aspect of ubiquitin-proteasome function. J. Biol. Chem., 279, 15427–15433.

References 109 Ando, H., Wen, Z.M., Kim, H.Y.,

110

111

112

113

114

115

116

Valencia, J.C., Costin, G.E., Watabe, H., Yasumoto, K., Niki, Y., Kondoh, H., Ichihashi, M., and Hearing, V.J. (2006) Intracellular composition of fatty acid affects the processing and function of tyrosinase through the ubiquitin– proteasome pathway. Biochem. J., 394, 43–50. Martinez-Esparza, M., JimenezCervantes, C., Beermann, F., Aparicio, P., Lozano, J.A., and Garcia-Borron, J.C. (1997) Transforming growth factor-beta 1 inhibits basal melanogenesis in B16/ F10 mouse melanoma cells by increasing the rate of degradation of tyrosinase and tyrosinase-related protein-1. J. Biol. Chem., 272, 3967–3972. Martinez-Esparza, M., JimenezCervantes, C., Solano, F., Lozano, J.A., and Garcia-Borron, J.C. (1998) Mechanisms of melanogenesis inhibition by tumor necrosis factoralpha in B16/F10 mouse melanoma cells. Eur. J. Biochem., 255, 139–146. Helenius, A., Marquardt, T., and Braakman, I. (1992) The endoplasmic reticulum as a protein-folding compartment. Trends Cell. Biol., 2, 227–231. Petrescu, S.M., Petrescu, A.J., Titu, H.N., Dwek, R.A., and Platt, F.M. (1997) Inhibition of N-glycan processing in B16 melanoma cells results in inactivation of tyrosinase but does not prevent its transport to the melanosome. J. Biol. Chem., 272, 15796–15803. Petrescu, S.M., Branza-Nichita, N., Negroiu, G., Petrescu, A.J., and Dwek, R.A. (2000) Tyrosinase and glycoprotein folding: roles of chaperones that recognize glycans. Biochemistry, 39, 5229–5237. Xu, Y., Bartido, S., Setaluri, V., Qin, J., Yang, G., and Houghton, A.N. (2001) Diverse roles of conserved asparaginelinked glycan sites on tyrosinase family glycoproteins. Exp. Cell. Res., 267, 115–125. Ando, H., Ichihashi, M., and Hearing, V.J. (2009) Role of the ubiquitin proteasome system in regulating skin

117

118

119

120

121

122

123

124

125

pigmentation. Int. J. Mol. Sci., 10, 4428–4434. Shigeta, Y., Imanaka, H., Ando, H., Ryu, A., Oku, N., Baba, N., and Makino, T. (2004) Skin whitening effect of linoleic acid is enhanced by liposomal formulations. Biol. Pharm. Bull., 27, 591–594. Galibert, M.D., Carreira, S., and Goding, C.R. (2001) The Usf-1 transcription factor is a novel target for the stressresponsive p38 kinase and mediates UV-induced tyrosinase expression. EMBO J., 20, 5022–5031. Bellei, B., Maresca, V., Flori, E., Pitisci, A., Larue, L., and Picardo, M. (2010) p38 regulates pigmentation via proteasomal degradation of tyrosinase. J. Biol. Chem., 285, 7288–7299. Smalley, K. and Eisen, T. (2000) The involvement of p38 mitogen-activated protein kinase in the alpha-melanocyte stimulating hormone (alpha-MSH)induced melanogenic and antiproliferative effects in B16 murine melanoma cells. FEBS Lett., 476, 198–202. Hall, A.M. and Orlow, S.J. (2005) Degradation of tyrosinase induced by phenylthiourea occurs following Golgi maturation. Pigment Cell Res., 18, 122–129. Kobayashi, T. and Hearing, V.J. (2007) Direct interaction of tyrosinase with Tyrp1 to form heterodimeric complexes in vivo. J. Cell. Sci., 120, 4261–4268. Park, H.Y., Perez, J.M., Laursen, R., Hara, M., and Gilchrest, B.A. (1999) Protein kinase C-beta activates tyrosinase by phosphorylating serine residues in its cytoplasmic domain. J. Biol. Chem., 274, 16470–16478. Iozumi, K., Hoganson, G.E., Pennella, R., Everett, M.A., and Fuller, B.B. (1993) Role of tyrosinase as the determinant of pigmentation in cultured human melanocytes. J. Invest. Dermatol., 100, 806–811. Bhatnagar, V., Anjaiah, S., Puri, N., Darshanam, B.N., and Ramaiah, A. (1993) pH of melanosomes of B 16 murine melanoma is acidic: its physiological importance in the regulation of melanin biosynthesis. Arch. Biochem. Biophys., 307, 183–192.

115

116

4 Biosynthesis of Melanins 126 Puri, N., Gardner, J.M., and Brilliant,

130 Martinez, J.H., Solano, F., Garcia-

M.H. (2000) Aberrant pH of melanosomes in pink-eyed dilution (p) mutant melanocytes. J. Invest. Dermatol., 115, 607–613. 127 Fuller, B.B., Spaulding, D.T., and Smith, D.R. (2001) Regulation of the catalytic activity of preexisting tyrosinase in black and Caucasian human melanocyte cell cultures. Exp. Cell Res., 262, 197–208. 128 Ancans, J., Tobin, D.J., Hoogduijn, M.J., Smit, N.P., Wakamatsu, K., and Thody, A.J. (2001) Melanosomal pH controls rate of melanogenesis, eumelanin/ phaeomelanin ratio and melanosome maturation in melanocytes and melanoma cells. Exp. Cell. Res., 268, 26–35. 129 Hearing, V.J. and Ekel, T.M. (1976) Mammalian tyrosinase. A comparison of tyrosine hydroxylation and melanin formation. Biochem. J., 157, 549–557.

Borron, J.C., Iborra, J.L., and Lozano, J.A. (1985) The involvement of histidine at the active site of Harding-Passey mouse melanoma tyrosinase. Biochem. Int., 11, 729–738. 131 Watabe, H., Valencia, J.C., Yasumoto, K., Kushimoto, T., Ando, H., Muller, J., Vieira, W.D., Mizoguchi, M., Appella, E., and Hearing, V.J. (2004) Regulation of tyrosinase processing and trafﬁcking by organellar pH and by proteasome activity. J. Biol. Chem., 279, 7971–7981. 132 Cheli, Y., Luciani, F., Khaled, M., Beuret, L., Bille, K., Gounon, P., Ortonne, J.P., Bertolotto, C., and Ballotti, R. (2009) αMSH and cyclic AMP elevating agents control melanosome pH through a protein kinase A-independent mechanism. J. Biol. Chem., 284, 18699–18706.

117

5 Inhibitors and Enhancers of Melanogenesis Alain Taïeb, Muriel Cario-André, Stefania Briganti, and Mauro Picardo

5.1 Introduction

In humans, normal levels of pigmentation differ among races, and reﬂect quantitative and qualitative differences in amount, size, and type of melanins synthesized and stored by melanosomes, which are speciﬁc melanin-containing lysosome-like intracellular organelles located inside melanocytes [1–3]. Each epidermal melanocyte is surrounded by approximately 36 keratinocytes, forming the so-called epidermal melanin unit, which plays a key role in the distribution of melanin pigments, but which is also under strong dermal inﬂuences [4, 5]. Phenotypic differences in constitutive pigmentation are not related to melanocyte numbers, which are almost the same in all skin types [6], but are determined by several other factors, including the level of melanogenic activity [7], the ratio of black/brown eumelanin to red/yellow pheomelanin [8, 9], melanosome transport, and distribution into neighboring keratinocytes [10] (see also Chapter 10). Thus, the melanin content and distribution inside the epidermal layers can be considered a sign of the healthiness of the skin. However, despite its photoprotective function in human skin, abnormal increased production and accumulation of melanin characterizes a large number of skin diseases, including acquired hyperpigmentation, such as melasma, postinﬂammatory melanoderma, freckles, ephelides, senile lentigines, and so on [11, 12]. On the contrary, a reduction of skin pigmentation is associated with degenerative processes, such as depigmentation in aged skin, which is considered a marker of melanocyte senescence, or with pathological conditions, such as vitiligo. The impact of hyper/hypopigmentation disorders is considerable across most human cultures and higher in general in dark-skinned populations, even for hyperpigmentation. The resulting psychosocial and cosmetic problems, and large proﬁtable market, have led to the more thorough investigation of melanogenic pathways and to the screening of the activity of a number of recognized and putative melanogenesis modulators.

Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

118

5 Inhibitors and Enhancers of Melanogenesis

5.1.1 Melanin Biochemistry 5.1.1.1 Melanin Biosynthesis Epidermal melanogenesis is a dynamic process, involving several regulating factors, which results in a peculiar composition of melanin pigments in accordance with skin type. Inside the melanosomes, the biosynthetic pathway for melanin formation is controlled by a combination of enzymatically catalyzed and chemical reactions. The ﬁrst and rate-limiting step of melanogenesis is the oxidation of the amino acid l-tyrosine to dopaquinone catalyzed by tyrosinase or polyphenoloxidase (EC 1.14.18.1, o-diphenol : O2 oxidoreductase). The enzyme uses molecular oxygen to catalyze the hydroxylation of the monophenol l-tyrosine to the o-diphenol 3,4-dihydroxyphenylalanine (dopa) and the oxidation of dopa to the o-quinone dopaquinone, which subsequently serves as precursor of both eumelanin and pheomelanin (see Chapters 3, 4, 6) [13, 14]. Dopaquinone is spontaneously converted to the monomeric indolic precursors (5,6-dihydroxyindole (DHI) and DHI2-carboxylic acid (DHICA)) of the black/brown pigment eumelanin. Moreover some other enzymes, like the tyrosinase-related protein (TRP-1/Tyrp1) and dopachrome tautomerase (Dct; also known as tyrosinase-related protein 2 (TRP-2/ Tyrp2)) may also play an important role in melanogenesis in vivo. TRP-2 catalyzes the tautomerization of dopachrome to DHICA, which is later oxidized to DHICAmelanin subunits. TRP-1 has been reported to catalyze the oxidation of DHICA to eumelanin. Subunits of eumelanin are composed by DHICA in conjunction with DHI, which is generated by the spontaneously lost carboxylic acid moiety of dopachrome. Moreover, TRP-1 is important for the correct trafﬁcking of tyrosinase to the melanosomes [15] and stabilization of its enzymatic activity [16, 17], and TRP-2 seems to be involved in the tyrosinase detoxiﬁcation inside melanosomes [18]. In the presence of thiol compounds, such as cysteine of glutathione, dopaquinone forms 2- or 5-S-cysteinyldopa that generates the benzothiazine precursors of the yellow/red soluble melanins, known as pheomelanins [19, 20]. In general, a mixed type of pheo- and eumelanin polymer is produced and deposited onto the melanosomal matrix proteins. 5.1.1.2 Tyrosinase Maturation and Degradation Tyrosinase contains six N-linked glycosylation sites that are conserved in human and mouse tyrosinases, and mutations of these critical protein maturation sites reduce tyrosinase catalytic function [21]. The altered ability to glycosylate tyrosinase induces the inhibition of its folding and maturation in the endoplasmic reticulum (ER) and Golgi, resulting in hypopigmentation [22]. Tyrosinase is degraded endogenously, at least in part, via the ubiquitin–proteasome system (UPS) [23], which selectively degrades intracellular ubiquitylated proteins, such as proteins misfolded in the ER and short-lived proteins [24–26]. Proteasome proteolyses tyrosinase via ER-associated protein degradation, a mechanism for quality control that involves retention in the ER and retro-translocation into the cytosol of misfolded or unassembled secretory proteins [27].

5.1 Introduction

5.1.1.3 Catalytic Site Tyrosinase is widely distributed throughout the phylogenetic scale from bacteria to mammals, and despite the high heterogeneity concerning the length and overall identity of the published sequences, its catalytic site is highly conserved among different species and shows high homology with TRP-1 and TRP-2 [28]. In particular, tyrosinase shows an active site consisting of a hydrophobic pocket, with a central copper-binding domain, which contains two peptidic segments, composed of strictly conserved amino acid residues, including three histidines [29]. Moreover, human tyrosinase shows the N-terminal signal peptide, which plays a relevant role in trafﬁcking and processing the enzyme, and the C-terminal hydrophobic transmembrane segment, which is rich in cysteine residues and is necessary for targeting tyrosinase to the melanosome [29–32]. The mechanism of catalysis of tyrosinase has been extensively investigated, due to its complexity and the peculiarity of the existence of two catalytic activities (tyrosinase hydroxylase and dopa oxidase) at the same active site [33, 34] (see Chapter 4). The link between copper ions and the three histidines is needed to effectively bind l-tyrosine to the active site and initiate melanin synthesis [35]. Thus, every alteration of peptide segments or copper ions can result in deregulation of tyrosinase activity. 5.1.2 Paracrine Signaling and Regulation of Epidermal Melanogenesis

Melanocytes work in close harmony with their neighboring cells in the epidermis. They are inﬂuenced by a variety of biological factors, including cytokines, growth factors, vitamins, and prostaglandins, which determine not only whether melanin is synthesized, but what type of melanin is produced. Presumably, these factors provide the complex signals that stimulate pigmentation after trauma, UV exposure, or other environmental stimuli that induce alterations in the levels of pigment production [36–38]. The expression of tyrosinase, related melanogenic enzymes, and melanosome structural proteins (MART-1 and PMEL17) may be modulated at the transcriptional level by the microphthalmia-associated transcription factor (MITF) [39, 40]. The binding of α-melanocyte-stimulating hormone (α-MSH) to the melanocortin 1 receptor (MC1R) activates adenyl cyclase and the subsequent release of cAMP, leading to activation of the protein kinase A (PKA) pathway. PKA induces the phosphorylation of cAMP-responsive element binding protein (CREB) transcription factors, which mediate MITF-M promoter activation to induce melanogenesis [41]. Interleukin-6 (IL-6) and the Wnt signaling pathway also regulate MITF [42–44]. Post-transcriptional MITF activation involves its transient phosphorylation via ribosomal S6 kinase (RSK), glycogen synthase kinase-3β (GSK3β ), p38 stress signaling, and mitogen-activated protein kinase (MAPK) pathways [45–47]. Moreover, paracrine linkages among keratinocytes, ﬁbroblasts, and melanocytes within the skin play important roles in regulating epidermal melanization. In response to various stimuli, human keratinocytes secrete various cytokines that serve as mitogens or melanogens for human melanocytes, including endothelin (ET)-1, stem cell factor (SCF), basic ﬁbroblast growth factor (bFGF), and α-MSH

119

120

5 Inhibitors and Enhancers of Melanogenesis

[48–51]. Overexpression of SCF and c-kit in the dermis has been proposed to play an important role in the mechanism of hyperpigmentation in melasma [52]. 5.1.3 Methods of Study

There has been great variation in results reported for different bioactive compounds targeted at regulating melanin production; in part, this variation derives from the diverse conditions of melanogenic assays employed, the different cell lines (including melanoma cells) used as targets, and the different endpoints used to establish efﬁcacy. In an attempt to standardize the assessment of novel bioactive agents for therapy of pigmentary lesions, a method called STOPR (standardized testing of pigmentary regulators) has been developed [53]. This protocol involves initial screening of putative melanogenic compounds using puriﬁed tyrosinase, followed by evaluation of cell proliferation, total melanin accumulation, and melanogenic potential using cultured pigmented murine melanocytes. However, even if the results obtained with the STOPR method are reproducible and economic, and can provide important insights into the mechanism(s) of depigmenting agents, more details about the effects toward human pigmentation process are needed. Up to now, in fact, the majority of in vitro testing studies have been performed by using mushroom tyrosinase, whereas the use of human tyrosinase can afford results more comparable to those observable in human skin [54]. To evaluate the active role of keratinocytes in melanogenesis modulation mammalian skin or keratinocyte–melanocyte cocultures should be preferred as testing models rather than only pure melanocyte cultures. For instance, arbutin is a more potent inhibitor of melanin formation on melanocytes cocultured with keratinocytes than on pure melanocyte cultures [55]. Furthermore, to analyze the contribution of dermal cells to the pigmentation process skin reconstructs including ﬁbroblasts should be used [5]. However, no experimental models are available for evaluating the effect of compounds able to induce hypopigmentation not only by inhibiting de novo melanin production, but also by increasing desquamation or bleaching pigment. Recently, new approaches have been proposed to discover new depigmenting agents. Chemical genetic screening performed in cultured Melan-A cells has been successfully employed to show depigmenting properties in vitro and shed new light on pathways inﬂuencing mammalian pigmentation [56, 57]. Potent pigmentation enhancers or inhibitors have been identiﬁed by screening tagged triazine-based combinatorial libraries in immortalized murine melanocytes, albino melanocytes, or zebraﬁsh. The mechanism of action of the identiﬁed compounds were investigated in zebraﬁsh and in skin equivalents, as useful tools to address the ability of these molecules to modulate pigmentation in a model more closely representative of human skin. In particular, zebraﬁsh may represent a useful and cost-effective alternative animal model, and can afford a good method to identify and characterize mutations that alter pigmentation or inﬂuence the development or subsequent migration of pigmentation precursor cells in the neural crest [58, 59].

5.2 Depigmenting Agents

5.2 Depigmenting Agents

Table 5.1 summarizes the agents discussed in the text according to the main pharmacological target and Table 5.2 indicates the major clinical indications. 5.2.1 Agents Acting Prior to Melanin Synthesis 5.2.1.1 Transcriptional Inhibition of Melanogenic Enzymes Considering the relevant role played by MITF in regulating melanogenesis, modulation of signaling pathways involved in its activity may have potential therapeutic use. Substances able to inhibit MITF expression and activity, as well as the extracellular signal-regulated kinase (ERK) and serine/threonine kinase (Akt)/protein kinase B (PKB) pathways could represent depigmenting agents [82, 83]. Among these, all-trans-retinoic acid (ATRA) is able to interfere with melanocyte development and melanogenesis. ATRA has a lightening effect on hyperpigmented lesions of photodamaged human skin [74, 75], but its depigmenting mechanism is not well known. This compound exerts controversial effects on melanocytes: it induces the transcription of tyrosinase by protein kinase C (PKC) activation and MITF expression, leading to melanocyte differentiation, and it promotes apoptosis of differentiated melanocytes via the caspase-3 pathway and Bcl-2 downmodulation [84]. In B16 melanoma cells, ATRA caused growth arrest and differentiation, and increased the level of PKC, which activates tyrosinase via phosphorylation of serine residues of cytoplasmic domain, thus enhancing melanin production [85]. Several researchers reported that ATRA and other vitamin A derivatives are able to increase pigmentation both in vitro and in animal models [86, 87]. However, other data in the literature suggest that ATRA activate melanogenesis only in cells or subjects with low amounts of melanin, whereas it signiﬁcantly improves the irregular hyperpigmentation associated with photoaging of inﬂammation [88]. Solar lentigo, a common component of photoaged skin, has been reported to represent an impaired homeostasis of the epidermal unit due to chronic sun exposure [89]. In the pathogenetic model suggested by the authors based on immunohistochemistry and electron microscopy, keratinocytes are more altered than melanocytes, suggesting that abnormal pigment retention in keratinocytes is the primary defect in solar lentigo. This evidence may partly explain the therapeutic effect of retinoids, which ﬁrstly targeted keratinocytes. Recently, ATRA and retinol have been found to inhibit cAMP-mediated melanin production in B16 melanoma cells and to downregulate tyrosinase, as well as TRP-1 expression prior to translation [90]. Although the role of PKC in the induction of melanogenesis remains controversial the existence of an interaction between the PKC pathway and the cAMPdependent PKA pathway has been suggested by the evidence that PKC inhibitors sphingosine and calphostine also inhibit the melanogenic response dependent on α-MSH [77]. The link between PKC- and cAMP-dependent pathways in regulating

121

Transcription inhibition 4-tertiary butylphenol

hydroquinone

4-hydroxyanisole N-acetyl-4-S cystaminylphenol monobenzyl ether of hydroquinone 4-n-butylresorcinol α-arbutin

C2-ceramide

zeolite

diosgenin

L. apetalum extract

dihydrolipoic acid

4-tertiary butylcathecol

Interference with tyrosinase activity

lysophosphatidic acid

protein kinase C inhibitors

all-trans-retinoic acid

During melanin synthesis

Classiﬁcation of depigmenting agents according to main pharmacological impact.

Before melanin synthesis

Table 5.1

reduction of melanosome transfer

inhibition of melanocytic dendrite formation

niacinamide

methylophiopogonanone B centaureidin

Inhibitors of melanosome transfer

After melanin synthesis

122

5 Inhibitors and Enhancers of Melanogenesis

Post-translational modiﬁcation of melanogenic enzymes

kojic acid gentisic acid and methyl ester of gentisic acid azelaic acid aloesin ellagic acid resveratrol oxyresveratrol

N-butyldeoxynojircimicin

deoxymannojirimycin

d-pantetheine-S-sulfonate

resveratrol

quinolines

linoleic acid

phospholipase D2

TRP-2 (Dct) modulation

deoxyarbutin

glucosamine

pyrroloquinoline quinone

all-trans-retinoic acid

adapalene

β-carotene

tarazotene

(Continued)

all-trans-retinoic acid

retinoids

2,6-dimethoxy-N-(4methoxyphenyl) benzamide

lectins neoglycoproteins

RWJ-50353 soybean extracts (soybean trypsin inhibitor and Bowman– Birk protease inhibitor)

Acceleration of epidermal turnover

inhibition of membrane glycosylation

inhibition of protease-activated receptor 2

After melanin synthesis

oxyresveratrol derivative

gnetol

arbutin

During melanin synthesis

tunicamycin

Before melanin synthesis

5.2 Depigmenting Agents 123

(Continued)

Before melanin synthesis

Table 5.1

peroxidase inhibitors

interference with the melanogenic pathway

interference with byproduct production

methimazole

N,N′-dilinoleylcystamine

cystamine

thioctic acid

gallic acid

pycnogenol

piceatannol

phytoncide

licorice extracts

salicylic acid

α-tocopherol ferulate

linoleic acid

lactic acid

α-tocopherol

6-hydroxy-3,4dihydrocumarins

glycolic acid

disodium isostearyl 2-O-lascorbyl phosphate

trichloroacetic acid

magnesium l-ascorbate-3-phosphate

chemical peels

retinaldehyde

After melanin synthesis

ascorbic acid

During melanin synthesis

124

5 Inhibitors and Enhancers of Melanogenesis

5.2 Depigmenting Agents Table 5.2

Clinical applications of depigmenting agents.

Melanogenic step

Treatment

Clinical indication

Main references

Interference with tyrosinase activity

hydroquinone

melasma

reviews: [60, 61] (Cochrane database)

postinﬂammatory hyperpigmentation solar lentigo 4-n-butylresorcinol

melasma

[62, 63]

solar lentigo deoxyarbutin

solar lentigo

[64, 65]

azelaic acid

melasma

reviews: [60, 61] (Cochrane database)

postinﬂammatory hyperpigmentation

Interference with byproducts production

ellagic acid + arbutin

melasma

aloesin

postinﬂammatory hyperpigmentation

aloesin + arbutin

postinﬂammatory hyperpigmentation

[67]

kojic acid

melasma

[68]

ascorbic acid

melasma

[69]

[66]

postinﬂammatory hyperpigmentation solar lentigo

Peroxidase inhibitors

α-tocopherol

melasma

thioctic acid

melasma

pycnogenol

melasma

methimazole

melasma

[70, 71]

postinﬂammatory hyperpigmentation Inhibitors of melanosome transfer

niacinamide

postinﬂammatory hyperpigmentation

[72, 73]

solar lentigo (Continued)

125

126

5 Inhibitors and Enhancers of Melanogenesis Table 5.2

(Continued)

Melanogenic step

Treatment

Clinical indication

Main references

Acceleration of epidermal turnover

all-trans-retinoic acid

melasma

reviews: [60, 61, 74, 75] (Cochrane database)

solar lentigo tarazotene

postinﬂammatory hyperpigmentation

[52, 76]

solar lentigo adapalene

melasma

[52, 74, 77]

solar lentigo β-carotene

melasma

[78]

trichloroacetic acid

melasma

reviews: [60, 61] (Cochrane database)

solar lentigo glycolic acid

melasma solar lentigo

salicylic acid

melasma

reviews: [60, 61] (Cochrane database) [79]

postinﬂammatory hyperpigmentation

Combined therapy

linoleic acid

melasma

[80]

licorice extracts

melasma

[81]

hydroquinone + alltrans-retinoic acid + a steroid

melasma

reviews: [60, 61] (Cochrane database)

postinﬂammatory hyperpigmentation solar lentigo

hydroquinone + alltrans-retinoic acid

melasma

reviews: [60, 61] (Cochrane database)

melanogenesis has been conﬁrmed by the evidence that MITF-M is a key factor for PKC-β [91]. Bisisolylmaleimide GF 109203X, a selective inhibitor of PKC, was found to reduce melanin production in human melanocytes, and to decrease both basal and UV-induced skin pigmentation in guinea, pigs, suggesting its possible application for the treatment of unwanted pigmentation [92]. Moreover, lysophosphatidic acid (LPA), an intercellular lipid mediator, which is known to have growth factor-like activity, contributes to reduced melanin synthesis via the downregulation of MITF [93]. Sphingomyelin is the most common sphingolipid in the skin

5.2 Depigmenting Agents

and ceramides are products of sphingomyelin hydrolysis by sphingomyelinases [94], suggesting that sphingolipid breakdown products play important roles in the regulation of epidermal proliferation, differentiation, and melanin synthesis. C2ceramide was found to inhibit cell proliferation and melanogenesis in human melanocytes, and in a mouse melanocyte cell line [83, 95, 96]. The suggested mechanism by which C2-ceramide decreases the pigmentation of melanocytes involves a sustained and long-lasting activation of ERK1/2 and Akt kinase, resulting in a reduction of MITF and inhibiting tyrosinase activity. Although rapid activation of ERK induces MITF phosphorylation and degradation, its delayed activation may lead to the inhibition of MITF expression Zeolites have crystalline open framework structures constructed from SiO4 and AlO4 tetrahedra linked through oxygen bridges. Each oxygen atom is shared by two silicon or aluminum atoms. Silicates and aluminosilicates possess biological activity and also are nontoxic agents. Zeolite inhibits MITF-mediated tyrosinase expression and melanin synthesis via increasing ERK activity in B16F10 melanoma cells [97]. Recently, heterocyclic pyrimidine compounds have been reported to protect epidermal cells from UV-B-induced damage and also to suppress melanogenesis via an MITF-mediated decrease of tyrosinase expression [98]. Compounds able to interfere with MITF post-transcriptional regulation can modulate the pigmentation process. Diosgenin, a steroidal saponin that is found in several plants, shows inhibitory effects on melanogenesis by activating phosphatidylinositol-3-kinase (PI3K) signaling. Akt and GSK3β are downstream molecules of PI3K-mediated signaling, and have been widely implicated in cell homeostasis by its ability to phosphorylate a broad range of substrates [99]. Diosgenin increases phosphorylated levels of both Akt and GSK3β, leading to a subsequent activation of MITF [80]. Cytokines reportedly regulate melanogenesis, and are major regulators of tyrosinase and related proteins. In Mel-Ab cells, transforming growth factor (TGF)-β1 inhibits pigment formation, and is able to interfere with tyrosine synthesis and possibly with the intracellular stability of the protein itself [100–102]. The proposed mechanism is a delayed ERK activation and subsequent reduction of MITF and tyrosinase production. In addition, IL-6 and tumor necrosis factor (TNF)-α are able to decrease pigmentation by acting on tyrosinase activity, although they exhibit an inhibitory effect in much higher concentrations [103, 104]. An extract of Lepidium apetalum (ELA) was reported to decrease UV-induced skin pigmentation in brown guinea pigs and melanogenesis of HM3KO human melanoma cells. The observed decrease of tyrosinase mRNA and protein expression as well as melanin content is proposed to be connected to ELA-mediated increase of IL-6 in keratinocytes, and the subsequent decreased expression of MITF [105]. The central role of MITF in melanogenesis is conﬁrmed by the evidence that some antioxidants, such as dihydrolipoic acid, α-lipoic acid (LA), and resveratrol, acts as depigmenting agents by a mechanism that does not involve the antioxidant activity per se. These compounds affect MITF and tyrosinase promoter activities, and reduce both endogenous and induced MITF mRNA and protein levels. Moreover, these agents induce depigmentation in melanocyte cultures, by counteracting

127

128

5 Inhibitors and Enhancers of Melanogenesis

the forskolin- and UV-B-stimulated promoter activities of these genes, and signiﬁcantly reduce tyrosinase activity [46]. Dihydrolipoic acid was found to depigment dark-skinned swine and to prevent UV-B-induced tanning in vivo. However, due to the fact that MITF is essential in regulating melanocyte differentiation, development, and survival [40, 106], to introduce compounds able to reduce MITF expression and activity as therapy for skin hyperpigmentation can lead to melanocyte death, as suggested by the decreased resistance to UV-induced apoptosis in melanocytes deﬁcient in MITF expression [107]. 5.2.1.2 Post-Translational Modiﬁcation of Melanogenic Enzymes The major post-translational modiﬁcation of melanogenic enzymes is asparaginelinked glycosylation, a process critical for the proper maturation of enzymes and the release of their soluble form [21, 108]. Melanogenic proteins show different glycosylation patterns, probably due to the different three-dimensional structures of tyrosinase, TRP-1, and TRP-2 [109]. Agents able to inhibit N-glycosylation result in improper protein folding and reduction of melanosome maturation, inhibition of melanosomal enzyme activities, or suppression of melanogenesis [110, 111]. In murine as well as in human melanoma cells, treatment with speciﬁc inhibitors of lipid carrier-dependent glycosylation of protein, such as tunicamycin or glucosamine, is able to decrease pigmentation and alter melanosome structure and biochemical functions [112, 113]. N-Butyldeoxynojircimicin inactivates tyrosinase and dramatically decreases melanin content in B16 melanoma cells, by inhibiting α-glycosidase I, an early step of N-glycosylation [114]. The same compound induced only a slight decrease of dopa oxidase activity of TRP-1, suggesting that this enzyme is able to bypass the glycosidase blockade. Recently, it was found that the pH-sensitive liposomes increased the glycosylationinhibiting and, thus, pigment-lightening effects of N-glycosylation inhibitors in vitro [115]. Treatment of HM3KO melanoma cells with deoxymannojirimycin, an inhibitor of α-1,2-mannosidase, which is thought to be responsible for late glycan processing, showed inhibition of glycosylation, transportation of tyrosinase to the melanosome and melanin synthesis [116]. In B16 melanoma cells, BMY-28565, a tyrosinase inhibitor, inhibited melanogenesis by depressing tyrosinase activity with no impact on tyrosinase mRNA levels and it has been suggested that this compound inhibited tyrosinase by modifying the sugar moieties of the enzyme [117]. Several compounds, including calcium d-pantetheine-S-sulfonate [118], ferritin [119], and glutathione [120], have been found to act as depigmenting agents by interfering with melanogenic enzyme glycosylation. Regarding calcium d-pantetheine-S-sulfonate, the similarity with the chemical structure of coenzyme A, which is involved in intracellular transport of protein, can play a relevant role in its mechanism of action [121]. Ferritin downmodulation and the subsequent intracellular stress conditions have been reported to substantially inﬂuence proper maturation of tyrosinase [119]. Resveratrol treatment of melanocytes determined depigmentation by increasing the retention inside of ER of immature tyrosinase and disrupting the trafﬁcking of the enzyme from the ER to the Golgi, leading to a dramatic decrease of functional tyrosinase [122].

5.2 Depigmenting Agents

Apart from the inhibition of tyrosinase glycosylation and the blockade of maturation and transfer processes of the enzyme, the bleaching effect of glutathione involves other mechanisms, such as chelation of copper ions at the active site, switch from eumelanogenesis to pheomelanogenesis, quenching of free radicals produced during melanogenesis, and modulation of depigmentation induced by melanocyte toxic compounds [54, 120]. Furthermore, intracellular pH is an important factor in the regulation of tyrosinase function, dramatically affecting tyrosinase enzymatic activity, and it is critical to the sorting of proteins to melanosomes [123, 124]. In Melan-A cells, quinoline analogs seem to disrupt the intracellular trafﬁcking of tyrosinase gene family proteins, probably with a mechanism involving their ability to modify pH of lysosomal compartment, leading to an alteration of subcellular localization of lysosome-associated membrane protein-1 and the subsequent reduction of pigmentation [125]. 5.2.1.3 Increased Tyrosinase Ubiquination Activation in vitro of tyrosinase degradation has been reported after treatment in vitro with different factors regulating the UPS, such as basic polypeptides, sodium dodecyl sulfate, guanidine-HCl, amino acids, or fatty acids [27, 126, 127]. Fatty acids, which are major components of cell membranes and are highly involved in intracellular signaling and in the binding of nuclear receptors [128], have been identiﬁed as intrinsic regulators of the UPS. There are several reports on the ability of fatty acid to induce hypopigmentation, by interfering at different steps of melanogenic pathway [129, 130]. Linoleic acid determined a reduction of melanin content without relevant changes of melanosome numbers or tyrosinase mRNA levels, suggesting a mechanism of action involving maturation and/or degradation of tyrosinase [131]. Fatty acids have been shown to have contrasting effects on the UPS, as unsaturated fatty acids, such as linoleic acid, increases the ubiquitination of tyrosinase, while palmitic acid decreases this process, leading to accelerated or decelerated degradation of tyrosinase by proteasomes, respectively [128, 129]. Topical application of linolenic, linoleic, and oleic acids (efﬁciency in decreasing order) has been shown to decrease UV-induced hyperpigmentation, suggesting that speciﬁc regulators of the UPS could be useful for the prevention and/or treatment of hyperpigmentary disorders. As with unsaturated fatty acids, phospholipase D2 also decreases melanogenesis through the same ubiquitin-mediated degradation of tyrosinase [132]. However, it has been also described that phospholipases and arachidonic acid produced a stimulation of melanogenesis [133], and it is possible that hyperpigmentation associated with the UV light response are mediated by those agents. If so, phospholipases and unsaturated fatty acids do not always exhibit hypopigmenting effects. 5.2.2 Agents Acting During Melanin Synthesis 5.2.2.1 Interference with Tyrosinase Due to its central role in regulating pigmentation, one of the most obvious cellular targets for depigmenting agents is the enzyme tyrosinase. Considering that the

129

130

5 Inhibitors and Enhancers of Melanogenesis

tyrosinase active site is characterized by a hydrophobic pocket into which the phenyl ring of the substrate is inserted, high stereo-speciﬁcity for l-isomers, and spatial restriction imposed by the distance between the phenyl ring and the groups on the side-chain, this suggests that a putative inhibitor should have a correspondent structure. Inhibition of tyrosinase activity can be obtained in different ways, including: (i) competitive inhibition, (ii) noncompetitive inhibition, and (iii) chelation of the copper atoms at the active site. Inhibitor strength is usually expressed as the inhibitory IC50 value, which is the concentration of an inhibitor needed to inhibit half of the enzyme activity in the tested condition. However, the classiﬁcation of tyrosinase inhibitors based on the structure and mechanism of action is not easy due to the large amounts of products reported and the controversial results of the kinetics performed employing different testing systems from mushroom tyrosinase, mammalian tyrosinase, melanocytic cultures, cocultures of keratinocytes and melanocytes, and ﬁnally in vivo application to animal skin. Furthermore, several compounds are able to inhibit tyrosinase activity with multiple mechanisms, such as interfering with residues of both catalytic and regulatory sites or being metabolized to a product able to act as a competitive as well as a noncompetitive inhibitor. A competitive inhibitor might be a copper chelator, nonmetabolizable analog, or derivative of the true substrate and is characterized by the capacity to bind the active site of the enzyme in a reversible manner, avoiding the interaction with the natural substrate and decreasing the enzyme–substrate afﬁnity, as demonstrated by the increase of Michaelis–Menten constant (Km). In contrast, an uncompetitive inhibitor can bind only to the enzyme–substrate complex. A mixed (competitive and uncompetitive mixed) type inhibitor can bind not only with the free enzyme, but also with the enzyme–substrate complex. For most mixed-type inhibitors, their equilibrium binding constants for the free enzyme and the enzyme–substrate complex, respectively, are different. However, a special case among the mixed inhibitors is the noncompetitive inhibitors, which bind to the free enzyme and an enzyme–substrate complex with the same equilibrium constant, inducing a decrease of enzymatic activity maximum velocity (Vm) and not modifying Km. As ﬁnal result both classes of inhibitors determined a reduction of tyrosinase kinetics and melanin synthesis. Compounds with electron donor groups may act as substrates and those with powerful electron acceptor groups may induce a competitive inhibition of tyrosinase. Inhibitors of tyrosinase may be grouped into two general categories. The ﬁrst includes compounds that possess an aromatic ring with functional groups, which form hydrogen bonds. The second category includes anions and molecules that form complexes with copper. In the ﬁrst category are included compounds containing either a 4-substituted phenol moiety or catechols, structurally similar to tyrosine or dopa, which, as alternative substrates of tyrosinase, are oxidized by the enzyme without producing melanin pigment [134, 135]. The 4-substituted phenol group has the ability to bind to the binuclear active site of the enzyme and inhibit its catalytic activity. The catechol structure, with two hydroxyl (OH) groups at the ortho positions, may behave as a chelator to the copper ions of tyrosinase. However, the

5.2 Depigmenting Agents

quinones generated can react with sulfydryl group of melanosomal proteins and/ or essential enzymes, interfering with cell growth and proliferation, and irritating the skin [134]. The ﬁrst compound introduced as a skin-lightening agent was hydroquinone (HQ), a phenolic compound chemically known as 1,4-dihydroxybenzene, which is considered one of the most effective inhibitor of melanogenesis in vitro and in vivo, and it has been deﬁned as the gold standard to compare the effectiveness of new depigmenting molecules. This compound has been widely used for the treatment of skin pigmentation disorders, such as melasma, or postinﬂammatory hyperpigmentation [61, 136–138]. Due to its structural homology with melanin precursors, HQ covalently binds to histidine or interacts with copper at the active site of tyrosinase, leading to the inhibition of the enzymatic oxidation of tyrosine and phenol oxidases. It also causes inhibition of RNA and DNA synthesis, and may alter melanosome formation and selectively damage melanocytes [139]. The decrease of melanin production is a direct consequence of HQ-induced suppression of melanocyte metabolic processes. Moreover, the ability of HQ to produce reactive oxygen species (ROS), inducing oxidative damage of membrane lipids and proteins, such as tyrosinase, and to deplete glutathione has been suggested to contribute in inhibiting pigmentation [54, 140–142]. Due to its effectiveness, the use of this molecule for whitening skin treatments was rapidly extended. The efﬁcacy and adverse effects of 4% HQ were evaluated in a double-blind placebocontrolled trial [143] and it has been reported that HQ is generally considered very safe [144]. However, due to its behavior as strong oxidant and its rapid transformation to quinones, which are melanocytotoxic compounds, HQ has been suggested to be toxic when applied on the skin in high concentration [145]. Commonly reported side-effects of HQ are skin irritation or contact dermatitis, which can be easily treated with topical steroids, but in a few patients, after a prolonged topical application of HQ in high concentration, permanent hypomelanosis or exogenous ochronosis, a quite irreversible hyperpigmentation in the treatment area, have been reported. To limit its cytotoxicity and side-effects the HQ dosage was reduced to 2%, a concentration that has been reported to improve hypermelanosis in 14– 70% of patients [146]. Moreover, it has been frequently used for many years, mixed with other compounds to increase its efﬁciency [61, 147, 148] in a number of melasma treatments. However, due to the hazard of long-term treatments, predisposing even to carcinogenicity [149], today the human use of HQ has important legal restrictions, and the addition of this compound in cosmetics has been banned by the European Commission (24 th Directive 2000/6/EC) and formulations containing HQ are available only by prescription of physicians or dermatologists. Several other phenolic compounds have been screened or used, and studies on the relationship between chemical structure and tyrosinase inhibitory activity have been performed. Methylation of one hydroxyl of para-hydroxylated monophenols favored the nucleophilic attack at the active site, enhancing the catalytic efﬁciency [150]. 4-Tertiary butylphenol has been reported to induce in human melanocytes a competitive inhibition of both tyrosine hydroxylase and dopa oxidase activities of tyrosinase without a cytotoxic response [138]. However, exposure to 4-tertiary

131

132

5 Inhibitors and Enhancers of Melanogenesis

butylphenol and 4-tertiary butylcathecol, which are used in the polymer industry, may cause leucoderma [151]. These substances generate reactive quinones and activate apoptotic process, leading to a loss of functional melanocytes [152]. Melanocytes from subjects with different phototypes demonstrated comparable sensitivity to 4-tertiary butylphenol, suggesting that the cytotoxic effect is completely independent from the tyrosinase activity and melanin content. Due to the presence of lypophilic groups, 4-hydroxyanisole (p-hydroxy-methoxybenzene) and N-acetyl-4-S-cystaminylphenol (4-SCAP) penetrate within the epidermis and target pigmented cells, showing depigmenting properties [142, 153–155]. 4-Hydroxyanisole is an alternative substrate for tyrosinase, being oxidized by the enzyme and in melanoma cell lines exhibits melanocytotoxic effects stronger than those exerted by HQ [156, 157]. Clinical effectiveness of 4-hydroxyanisole in skin hyperpigmentation has been demonstrated in combination with treinoin in solar lentiges and related hyperpigmented lesions [158]. Topical treatment with 4-SCAP of Yucatan pigs induces a stable but reversible depigmentation with less irritant effects as compared to HQ [159]. The activity seems to be related to: (i) a marked decrease in the number of functioning melanocytes and melanized melanosomes, (ii) a decreased number of melanosomes transferred to keratinocytes, and (iii) a selective degeneration of melanocytes and deposition of melanin-like material in the Golgi apparatus, where tyrosinase is located [160]. Like HQ, the monobenzyl ether of HQ (MBEH) belongs to the phenol/catechol class of chemical agents. However, this molecule causes a permanent depigmentation of the skin by its ability to be metabolized inside cells to a reactive quinone, which is highly cytotoxic [140]. This property makes MBEH suitable for eliminating residual areas of normally pigmented skin in patients with diffuse vitiligo [161]. In general, all diphenols leading to a reactive quinone are putative cytotoxic agents. This type of hypopigmenting agent does so by its cytotoxic action on melanocytes rather than by tyrosinase inhibition. This is the case of 4-(phydroxyphenyl)-2-butanone [162] and 4-n-butylresorcinol [163, 164]. Moreover, 4-nbutylresorcinol has been shown to inhibit the activity of both tyrosinase and TRP-1, and to be a molecule well tolerated and signiﬁcantly effective in the treatment of melasma [62, 63, 165]. 4-Hydroxyphenyl α-d-glucopyranoside (α-arbutin) is a naturally occurring HQ β-d-gluconopyranoside, which is enzymatically synthesized from HQ and saccharides [166, 167]. This compound shares with HQ the ability of acting as an alternative substrate of mammalian tyrosinase at noncytotoxic concentrations [168] and its inhibitory activity has been tested against tyrosinase from different sources, such as mushroom, B16 mouse melanoma, and human melanoma cells [169]. The treatment with α-arbutin determined a reduction of melanin synthesis also in human epidermal equivalents containing melanocytes [170]. Interestingly, in the medium of this experimental model about 70% of the applied α-arbutin was recovered, but HQ was not detected, suggesting that the activity of α-arbutin is not due to HQ release. In addition, arbutin, a natural β-glycoside optical isomer of α-arbutin, usually found in cranberries, blueberries, wheat, and pears [171], is a good inhibitor of tyrosinase, without affecting expression and synthesis of the

5.2 Depigmenting Agents

enzyme in human melanocyte cultures [54, 172]. Moreover, this molecule inﬂuences DHICA polymerase activity and PMEL17/silver protein, and exerts an inhibitory effect on the maturation of melanosomes [172]. However, the depigmenting effect of arbutin has not been conﬁrmed in clinical trials [173] and its mild effectiveness has been attributed to a controlled release of HQ 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 [141]. To increase its efﬁciency, α-glucosides of arbutin have been chemically synthesized [174], possibly since they are easier hydrolyzed to release HQ due the higher availability of α-glycosidases. Recently, deoxyarbutin, synthesized by removing every hydroxyl group of arbutin, has been identiﬁed as an excellent tyrosinase inhibitor in the screening of a number of candidate compounds [64]. In a hairless, pigmented guinea pig model and in human skin deoxyarbutin induced reversible inhibition of in situ tyrosinase activity, and rapid and sustained skin-lightening. This compound, as well as its derivatives, has been found to act as an effective competitive inhibitor of tyrosinase, leading to decreased melanin content in melanocytes [175]. This depigmenting effect is completely reversible, indicating no permanent suppression of tyrosinase activity and melanin synthesis. The presence of hydrophobic and less bulky groups at the para position of phenol increases binding to tyrosinase and decreases the potential for oxidation, leading to effective inhibition and reducing cytotoxicity [65, 176]. Deoxyarbutin and its derivatives can enter viable melanocytes at nontoxic concentrations, blocking enzyme activity of pre-existing tyrosinase, more than 90% of which is bound to the melanosome membrane, determining a post-translational regulation that can be reverted after the compounds are removed [54, 175]. Due to the presence of two copper ions at the active site of the enzyme, chelating compounds can be considered speciﬁc for tyrosinase. Kojic acid (5-hydroxy-2hydroxymethyl-4H-pyran-4-one) is an antibiotic produced by many species of Aspergillus and Penicillium with a high capacity for chelating transition metal ions such as Fe3+ and Cu2+. Kojic acid is a good tyrosinase inhibitor, but it has some undesirable side-effects. It may cause allergy [177] and it also has been related so some hepatic tumors in heterozygous mice deﬁcient in p53 [178]. The depigmenting action of kojic acid is attributed to its chelator ability [68], even if it could also interfere at different steps of melanin synthesis [179] and inhibit NF-κB activation in keratinocytes, contrasting the hyperpigmentation associated with inﬂammatory response [180]. Although in monotherapy kojic acid showed only a modest effectiveness, clinical trials have reported a skin-lightening effect of this compound in combination with other agents [60, 181]. Recently, some stable derivatives have been synthesized having more efﬁciency because their penetration through the skin is increased [182–184]. In this group, the most important ones are a derivative synthesized by joining two pyrone rings through and ethylene linkage [185] and kojyl-APPA (5-((3- aminopropyl)-phosphino-oxy)-2-(hydroxymethyl)-4H-1-pyran-4one), tested in melanoma cells and normal human melanocytes [186]. Gentisic acid (2,5-dihydrobenzoic acid; GA), a component of gentiana roots, showed good melanogenesis inhibition properties [187] and it has been explored

133

134

5 Inhibitors and Enhancers of Melanogenesis

for use in topical cutaneous applications. The methyl ester of GA (MG) has been found to inhibit mammalian tyrosinase in pure form, in crude cell-free extract, in addition to inhibition of melanin formation in cultured melanocytes. MG and GA are structurally similar to HQ, and it has been postulated that MG acts as a prodrug and inhibits tyrosinase activity following the liberation of HQ. Kinetics studies, however, have found a direct effect of MG on enzyme function, suggesting a binding of the copper ions in the active site, possibly in the presence of oxygen. MG can enter viable melanocytes at sufﬁcient concentrations to block tyrosinase activity, and thus de novo melanin biosynthesis, and it does not appear to inhibit TRP-2 or TRP-1 at concentrations higher than needed to inhibit tyrosinase [173]. MG was also less cytotoxic and mutagenic than HQ, although some low adverse side-effects were reported. Thus, this compound is proposed as a good candidate for skin-lightening, even if a test in skin models is needed to postulate its effectiveness in vivo. Compounds able to bind either amino or carbonyl groups may block the access of tyrosine to the active site, behaving as competitive inhibitors. Among these, azelaic acid (AZA), a naturally occurring 9-carbon dicarboxylic acid, derived from cultures of Pityrosporum ovale, is included [188, 189]. The depigmenting activity of AZA is mediated by inhibition of mitochondrial oxidoreductase activity and interference with DNA synthesis as well as competitive and reversible inhibition of tyrosinase activity in vitro (Ki = 2.73 × 10−3 M) [190–192]. Its lightening effect appears to be selective and most apparent in highly active melanocytes, with minimal effects in normally pigmented skin [193]. AZA has been used to treat melasma and postinﬂammatory hyperpigmentation [72, 194], and to arrest the progression of lentigo maligna to melanoma [195]. Topical AZA 15–20% has been reported to be as efﬁcacious as HQ improving melasma skin [196], even if not all authors agree with these therapeutic effects. Moreover, this compound has been reported to produce clinical and histological resolution in facial lentigo maligna, and to induce good results in the treatment of rosacea, solar keratosis, and hyperpigmentation associated with burns and herpes labialis. Furthermore, due to its mild and transient side-effects (i.e., erythema and cutaneous irritation) AZA is generally well-tolerated and can be used for extended periods [197]. Flavonoids are plant polyphenols classiﬁed as benzo-γ-pyrane derivatives, due to the presence of phenolic and pyrane rings in their chemical structures. These compounds include more than 4000 members identiﬁed in leaves, bark, and ﬂowers. Several natural medical treatments contain these polyphenols, due to their beneﬁcial effects, including anticancer, antiviral, and anti-inﬂammatory protection against UV damage [188]. Some of the therapeutic effects are related to the antioxidant and ROS scavenger properties of ﬂavonoids [198]. Moreover, their ability to chelate metals at the active site of metalloenzymes can explain melanin reduction induced by these compounds, even if their action at the distal part of the melanogenesis oxidative pathway can contribute to the hypopigmenting effect. The chemical structure of ﬂavonoid groups can inﬂuence their mechanism of action: ﬂavonoids with a α-keto group are similar to dopa and thus they show potent tyrosinase inhibition [188]; the presence of a free 3-hydroxy group improves

5.2 Depigmenting Agents

the ﬂavonoid’s capability to chelate copper in tyrosinase’s active site [199]. Aloesin, a natural hydroxymethyl chromone compound isolated from aloe vera, was demonstrated to be a competitive inhibitor of tyrosinase [200] and showed a dosedependent depigmenting effect in melanocytes alone in vitro. Due to the hydrophilic nature of aloesin and low penetration rate across skin layers, combined treatment with this compound and arbutin has been evaluated to assess the synergistic effects on tyrosinase activity. The two compounds act in a synergistic manner and interfere with tyrosinase function by different mechanisms: aloesin exhibits noncompetitive inhibition, while arbutin inhibits competitively [67, 201]. In pigmented human skin equivalents, aloesin showed dose-dependent reductions in tyrosinase activity and melanin content, and demonstrated a good safety proﬁle [202]. Considering that aloesin revealed no cytotoxicity, it has been suggested as a good alternative to HQ [142]. Ellagic acid is a naturally occurring polyphenol widely distributed in plants, and it has been reported to have anticarcinogenic, antiﬁbrosis, and antioxidative properties. This compound shows high afﬁnity to copper at the active site of tyrosinase and inhibits its activity by binding to the copper [203]. Furthermore, in brownish guinea pigs, as well as in human skin, topically applied ellagic acid induced a reversible inhibition of melanin synthesis only in UV-activated melanocytes [204]. Moreover, oral administration to humans of ellagic acid-rich pomegranate extract has a protective effect on slight sunburn caused by UV irradiation [205]. A recent study compared the effectiveness of gel formulations containing arbutin, synthetic ellagic acid, and plant extracts that contain ellagic acid on patients with melasma. The formulation prepared with plant extracts was suggested as an alternative to mono- or combined treatment of melasma, because it is nontoxic and has an effect of tyrosinase enzyme inhibition as demonstrated experimentally, as well as having antioxidant and UV-preventive effects [66]. Most of the hydroxystilbene compounds, like resveratrol (3,4,5-trihydroxystilbene), are derived from natural products found in herbal medicines and showed efﬁcient skin-lightening properties, due to their high afﬁnity for tyrosinase [206]. Apart from resveratrol and its derivatives, such as oxyresveratrol or gnetol, this group includes methoxylated or glucosylated derivatives (piceid-glucoside, rhapontigenin, and rhaponticin) with a structural skeleton comprised of two aromatic rings linked by an ethylene bridge [207, 208]. The number and position of hydroxyl substituents of hydroxystilbene compounds seem to play an important role on the inhibition of tyrosinase activity [206]. Kinetics studies revealed that oxyresveratrol and gnetol are more efﬁcient tyrosinase inhibitors than resveratrol [207]. Also, the trans-oleﬁn structure of the stilbene skeleton plays a relevant role in inhibiting tyrosinase, as suggested by the fact that trans-resveratrol is much more potent than the cis isomer [208]. In all cases, the inhibition of the enzyme is reversible and in vivo treatments should maintain high intracellular levels of the hydroxylated stilbene inside melanocytes. Some data indicated the ability of resveratrol to reduce MITF and tyrosinase promoter activation in B16 mouse melanoma can contribute to its effect on pigmentation [46, 54]. However, such evidence is controversial due to other data suggesting that resveratrol-treated human melanocytes displayed a

135

136

5 Inhibitors and Enhancers of Melanogenesis

steady-state tyrosinase RNA and consequently regulation of enzyme transcription did not induce depigmentation [122]. Additionally, further analysis of the resveratrol-treated normal human melanocytes showed ER-retained immature tyrosinase, suggesting disrupted trafﬁcking of tyrosinase in the Golgi–ER– lysosome and elevated proteolytic degradation [122]. In a study with 285 different plant extracts from oriental herbs, the best one seemed to be the extract from Morus alba L., which contains 2-oxyresveratrol. The extract inhibited tyrosinase activity remarkably, and showed no toxicity and no skin irritation effect [209]. 5.2.2.2 TRP-2 Modulation TRP-2 (Dct) is another major enzyme correlating with the melanogenesis pathway, even if it role in controlling melanin biosynthesis is not as crucial as the function of tyrosinase. This enzyme catalyzes the transformation of dopachrome into DHICA, acting as a “dopachrome conversion” factor [210]. In the melanization pathway, the conversion of dopachrome is a turning point that determines if DHIeumelanin or DHICA-eumelanin will be produced. Spontaneous conversion of dopachrome to DHI is faster than TRP-2-induced generation of DHICA, thus the main pathway is generally the production of DHI-eumelanin [211]. However, when generated, DHICA-eumelanin is able to inhibit the production of DHI-eumelanin. Due to the fact that DHICA-eumelanin pigment has a yellow/light brown color, which is milder than the light brown/black of DHI-eumelanin, the stimulation of TRP-2 is consequently described as having lightening effects. An oxyresveratrol derivative showed depigmenting properties by activating TRP-2 [212]. 2,6-DimethoxyN-(4-methoxyphenyl)benzamide, which was synthesized using a combination of benzoic acid and aniline, has been reported to accelerate dopachrome transformation into DHICA in the presence of TRP-2 and to exhibit signiﬁcant depigmentation ability on the UV-B-induced hyperpigmentation of brown guinea pig skin [213]. Compounds able to inhibit tyrosinase activity without interfering with TRP-2 functionality can also result in DHICA-eumelanin accumulation, leading to an improvement of skin hyperpigmentation. ATRA and pyrroloquinoline quinone, which functions as a ROS scavenger and protects cells against injury and DNA fragmentation, increased the generation of DHICA-eumelanin, even in the absence of a direct activation of TRP-2 activity, resulting in a skin-lightening effect [90, 214]. 5.2.2.3 Interference with Byproduct Production (Antioxidant and Reducing Agents) Skin exposure to sunlight generates ROS, which are able to stimulate epidermal melanogenesis either directly, acting as second messengers, or indirectly, through the induction of the production of cytokines by epidermal and dermal cells [215]. Compounds with redox properties can interfere with pigmentation process by neutralizing highly reactive o-quinone intermediates, and thus avoiding the oxidative polymerization during melanin synthesis, or by scavenging ROS. Ascorbic acid (AsA) interferes with the different steps of melanization, by reducing dopachinone and by blocking DHICA oxidation [140]. Moreover, AsA is highly unstable,

5.2 Depigmenting Agents

being quickly oxidized and decomposed in aqueous solution, and the degree of penetration into the skin is low, due to its prevalent hydrophilic nature. Different esters of AsA, such as magnesium l-ascorbate-3-phosphate (MAP), which is stable in water and in alkaline medium, have been used. MAP is absorbed percutaneously, reaches the epidermal layer, where can be hydrolyzed in AsA, and induces a lightening effect in both normal and hyperactive melanocytes [216]. Application of 10% MAP cream was shown to suppress melanin formation. A signiﬁcant lightening effect was seen clinically in 19 of 34 patients with melasma and solar lentigo. Furthermore, MAP has been shown to have a protective effect against skin damage induced by UV-B irradiation, by reducing, regardless of the drug administration route, lipid peroxidation and inﬂammation reaction in hairless mice [216, 217]. In vitro studies showed that MAP was converted to AsA as it crossed the epidermis, but that sodium ascorbate (AS-Na) did not pass through the epidermis. Furthermore, MAP was also converted to AsA in serum. These results suggest that the protective effect of MAP on UV-B-induced cutaneous damage is due to the conversion of MAP to AsA [218]. A novel amphiphilic ascorbic derivative, disodium isostearyl 2-O-lascorbyl phosphate (VCP-IS-2Na), signiﬁcantly suppressed the cellular tyrosinase activity of melanoma cells and melanocytes [219]. Treatment of human skin models with 1.0% VCP-IS-2Na decreased melanin synthesis, without inhibiting cell viability. A recent study compared both in vitro and in vivo the depigmenting effect of AsA or multivitamin [69]. In the melanoma cells stimulated by α-MSH multivitamin, which is composed by various vitamins, decreased melanin content more than AsA did. Multivitamin also revealed skin-lightening action as effective as AsA, clinically. The combination of hypopigmenting vitamins has an additive antimelanogenic effect with low cytotoxicity and downregulation of MITF, which gives enhanced efﬁcacy. In vitro, vitamin E (α-tocopherol) derivatives induce the inhibition of melanogenesis in epidermal melanocytes and show a potent inhibitory effect on tyrosinase [54]. The ability of α-tocopherol to act as a scavenger of ROS and chain-breaking antioxidant, interfering with lipid peroxidation of the melanocyte membrane, and to increase the intracellular glutathione content could explain its depigmenting effect [140]. However, during the antioxidant reaction α-tocopherol undergoes auto-oxidative processes and produces quinone radicals, leading to a loss of its antioxidative effect. The activation of the antioxidant recycling mechanism in which AsA, through the transfer of one electron, is able to regenerate α-tocopherol from its radicalic form, results in the more effective and long-lasting antioxidant response observed for coadministration of these compounds. In vivo, topical application of α-tocopherol and AsA reduced the tanning response, through an inhibition of UV-induced melanogenesis and proliferation of melanocytes. α-Tocopherol ferulate is a related compound in which α-tocopherol is linked by an ester bond to ferulic acid, an antioxidant that stabilizes α-tocopherol [220]. This molecule has been reported to inhibit melanin synthesis in normal human melanocytes, without interfering with the tyrosinase pathway. Moreover, in an attempt to provide an efﬁcient penetration into the skin, a formulation in which α-tocopherol ferulate is incorporated in liposomes has been synthesized [221]. The association between

137

138

5 Inhibitors and Enhancers of Melanogenesis

the protective effect of ferulic acid against α-tocopherol oxidation and the good percutaneous penetration of the liposomal preparation leads to the hypothesis that α-tocopherol ferulate added to liposomes may exert its lightening effects in active melanocytes providing an efﬁcient intracellular distribution of α-tocopherol. A study investigating the biochemical effect of α-tocopherol ferulate in human melanoma cells suggests the whitening effect is due to tyrosinase inhibition at the post-transcriptional level, possibly by an unidentiﬁed secondary molecule [142]. Tocopherols conjugated with phenols at position 4, such as resorcinol, seem to have a strong effect [222]. These compounds could exert a double action – the inhibition of tyrosinase by the phenol moiety and the lipophilicity and ROS scavenger activity of the α-tocopherol moiety. 6-Hydroxy-3,4-dihydrocumarins, which are antioxidants with α-tocopherol-like chemical structures [223], have been reported to have antimelanogenic activity in cultured normal human melanocytes at noncytotoxic concentrations and without interfering with tyrosinase activity. A possible mechanism is the acceleration of the biosynthesis of glutathione (GSH) within melanocytes, leading to the inhibition of tyrosine transfer to premelanosomes [224]. In an effort to ﬁnd new skin-lightening agents, natural plant extracts that contain compounds that have antioxidant and free-radical scavenger properties have been tested. Several studies identiﬁed substances, such as phytoncide [225], piceatannol [226], pycnogenol [227], and gallic acid [228], that exert antimelanogenic activity via their antioxidative property and/or their ability to suppress ROS while increasing the ratio between glutathione and oxidized glutathione (GSSG). LA (also known as thioctic acid), a disulﬁde derivative of octanoic acid, exhibits several biological effects, which include the quenching of ROS, metal chelation, interaction and regeneration of other antioxidants, redox regulation of protein thiol groups, and effects on gene expression and apoptosis [229, 230]. Lipoamide dehydrogenase, located exclusively in mitochondria, reduces free LA in dihydrolipoic acid, which is a potent antioxidant, being able to reduce ubiquinone and oxidized glutathione. LA has been reported to prevent UV-induced photo-oxidative damage, mainly through the downmodulation of NF-κB activation, and to inhibit tyrosinase activity, probably by chelating the copper ions at the active site [46, 231]. Recently, LA has been conjugated to poly(ethylene glycol) (PEG) of molecular weight 2000 (LA-PEG 2000) and its effects on melanogenesis has been evaluated [232] LA-PEG 2000 is a highly water-soluble molecule, which has lower cell cytotoxicity than LA. This compound signiﬁcantly suppressed the biosynthesis of melanin, and reduced both the activity and expression of tyrosinase in B16F10 melanoma cells. 5.2.2.4 Interference with the Melanogenic Pathway Reaction of thiol compounds with quinone intermediates of melanin can switch pigment synthesis from eumelanin to pheomelanin [233]. Elevated amounts of thiols, including glutathione, have been associated, in fact, with a prevalence of pheomelanin, leading to a lack of pigmentation observable both in humans and in mice [234]. Cystamine has been reported to be effective in reducing epidermal pigmentation in combination with HQ [147]. Cystamine as well as cysteamine, its

5.2 Depigmenting Agents

reduced form, can shift the melanin biosynthetic pathway in favor of pheomelanin by nucleophilic addition to dopaquinone, which is much faster than conversion of dopaquinone to dopachrome [235]. N,N′-dilinoleylcystamine, a compound of cystamine and linoleic acid connected by an ester bond, has a pigment-lightening effect on HM3KO melanoma cells by reducing the level of eumelanin and increasing the level of phaeomelanin [236]. 5.2.2.5 Peroxidase Inhibitors In the 1970s, Okun, Edelstein, Or, Hamada, and Donnellan suggested that peroxidase, an iron-containing enzyme, alone or in conjunction with tyrosinase catalyzes the oxidation of l-tyrosine to melanins [237, 238]. Hydrogen peroxide (H2O2) is produced as by product of DHI and DHICA oxidation, and the involvement of peroxidase–H2O2 systems in later stages of melanogenesis, during the oxidation of indolequinone precursors of eumelanin and benzothiazinylalanine precursors of pheomelanin, has been suggested [239, 240]. Recent studies have identiﬁed new physiological functions of tyrosinase, namely catalase (conversion of H2O2 to ½O2 and H2O) and peroxygenase (H2O2-dependent oxygenation of substrates), leading to scavenging of tyrosyl radicals, which are toxic oxidants of melanocytes. The role of H2O2 in melanogenesis has not been clearly deﬁned, with some reports indicating it functions to enhance pigment formation by regulating levels of tyrosinase [241], others suggesting the molecule serves as a potent inhibitor of tyrosinase [242]. Moreover, H2O2 is released by cytokines, such as TNF-α or TGF-β, which induce depigmentation [243–245], and it has been reported to produce a transient reduction of tyrosinase and other melanogenic proteins, through the downregulation of the MITF transcription factor [246]. The implication of H2O2 in the ﬁrst steps of melanogenesis has been supported by a study showing that H2O2 was generated during the oxidation of dopa to dopaquinone and that tyrosinasemediated diphenolase activity was enhanced by the endogenously generated H2O2 [247]. These data afford a possible explanation of depigmentation and reduction of the polymerization rate of eumelanin induced by inhibition of peroxidase [248]. Methimazole, an antithyroid agent belonging to the thionamide group, exerts inhibitory action towards both mushroom tyrosinase and peroxidase, and in brown guinea pigs induces mild-to-moderate inhibition of melanization, with morphologic changes of melanocytes [249]. This molecule showed whitening action also in humans and it has been suggested as a safe skin-depigmenting compound for topical treatment of hyperpigmentary disorders [70, 71]. 5.2.3 Agents Acting After Melanin Synthesis 5.2.3.1 Inhibitors of Melanosome Transfer A new approach to achieve depigmentation is to study the possible application as depigmenting agents of molecules able to interfere with melanosome maturation and transfer. Reduction of skin pigmentation can be achieved, in fact, also by decreasing melanosomal transfer from melanocytes to surrounding keratinocytes

139

140

5 Inhibitors and Enhancers of Melanogenesis

and subsequent melanosome processing inside recipient cells [250]. Several compounds have been found to inhibit melanosome transfer by interfering with different steps of this process. 5.2.3.1.1 Inhibition of Melanocytic Dendrite Formation To effectively transfer melanosomes, melanocytes should have a good dendrite generation and extension towards surrounding keratinocytes. The reorganization of the melanocyte cytoskeletal elements is needed for the extension of melanocyte dendrites [251]. Agents able to activate Rho, a small GTPase, which is involved in cell morphology and dendrite formation, can interfere with melanosome transfer. In melanocyte and keratinocyte cocultures, methylophiopogonanone B (5,7-dihydroxy-6,8-dimethyl-3-(4-methoxybenzyl)chroman-4-one) and centaureidin (5,7,3′-trihydroxy-3,6,4′-trimethoxyﬂavone), a ﬂavonoid glucoside, have been reported to activate Rho, inducing cytoskeleton disorganization, and to reduce melanosome transfer, by determining a reversible dendrite retraction [251–253]. These compounds did not inﬂuence melanin synthesis or the expression of melanogenic enzymes and can be considered as promising skin-lightening compounds, even if more studies are needed to verify their applicability in vivo. 5.2.3.1.2 Reduction of Melanosome Transfer Niacinamide, a biologically active form of niacin, is a precursor to the cofactors nicotinamide adenine dinucleotide and nicotinamide adenine dinucleotide phosphate, which participate in numerous enzymatic reactions [73, 201]. This compound has been proposed to have several actions in the skin, including anti-inﬂammation, prevention of photo-immunosuppression, and increased intercellular lipid synthesis [254]. Topical niacinamide is described to improve the appearance of photoaged facial skin (including texture, hyperpigmentation, redness, ﬁne lines, and wrinkles) [255–257]. Moreover, this molecule was found to inhibit melanosome transfer to keratinocytes both in vitro and in humans. However, the associated mechanism of action remains to be elucidated [73]. 5.2.3.1.3 Inhibition of PAR-2 protease-activated receptor 2 (PAR-2) is G-protein-coupled receptor and is activated by serine proteases, including trypsin or mast cell tryptase [258, 259]. Due to its expression in keratinocytes, but not in melanocytes, PAR-2 is involved in the modulation of skin pigmentation through the interaction between melanocytes and keratinocytes [260–262]. The inhibition of serine protease results in an impaired activation of PAR-2 on the keratinocyte leading to the accumulation of melanosomes within the melanocyte, and blocking the melanosomal transfer and melanin dispersion inside keratinocytes, subsequently inducing skin-lightening [263–265]. In epidermal reconstructs, RWJ-50353, a serine protease inhibitor, has been reported to induce accumulation of melanosomes in melanocytes and a concomitant negative feedback mechanism that slows pigment production [259].

5.2 Depigmenting Agents

Moreover, Yucatan swine skin treated with RWJ-50353 for an 8-week period demonstrated a dose-dependent, reversible skin-lightening effect [259]. Soybean extracts have been explored as depigmenting agents. Kunitz-type trypsin inhibitor (soybean trypsin inhibitor (STI)) and the Bowman–Birk protease inhibitor (BBI), two protein proteinase inhibitors isolated from soybean seeds, have been reported to reduce pigmentation in dark-skinned Yucatan swine, probably by inhibiting PAR-2 activation, leading to a reduction in keratinocyte phagocytosis and melanin distribution [265]. Other components of soybean can also play a role in the mechanism of action of STI and BBI: free fatty acids and their acyl-coenzyme A esters can inhibit trypsin and participate in PAR-2 inhibition; isoﬂavones may reduce dopa oxidase activity of tyrosinase; and phospholipids can assist the epidermal delivery of STI and BBI. 5.2.3.1.4 Inhibition of Membrane Glycosylation To transfer melanosomes, recognition between melanocytes and keratinocytes is needed. Due to their inﬂuence in intracellular trafﬁcking, endocytosis, and cell– cell recognition, lectins and neoglycoproteins have been investigated as compounds involved in this phenomenon. In an in vitro model, the role of glycosylated residues on melanocyte and keratinocyte membranes as part of receptor-mediated endocytosis facilitating melanosome transfer has been identiﬁed [250]. In the same paper, the authors suggest that lectins and neoglycoproteins play an inhibitory role in this process. In keratinocyte–melanocyte cocultures lectins were found to induce a reversible inhibition of melanosome transfer and this effect was enhanced by the presence of niacinamide [254]. 5.2.3.2 Acceleration of Epidermal Turnover Skin-lightening can also result from an increased rate of epidermal layer renewal and removal of pigmented upper layer keratinocytes. Thus, compounds able to activate skin desquamation can effectively act as depigmenting agents [140, 266]. 5.2.3.2.1 Retinoids Due to their capacity of inducing dispersion of keratinocyte pigment granules, interference with pigment transfer, acceleration of epidermal turnover, reduced cohesiveness of corneocytes, and desquamation [54, 267, 268], retinoids are considered hypopigmenting compounds. ATRA shows a depigmenting effect by inhibiting tyrosinase and TRP-1 transcription [90], and it has been reported to improve melasma, with some side-effects, such as erythema, peeling at the site of application, and postinﬂammatory pigmentation [54, 269–271]. This molecule is also used in combination in topical creams, such as a widely prescribed formulation proposed by Kligman and Willis containing 5% HQ, 0.1% tretinoin, and 0.1% dexamethasone [266]. However, ATRA requires long-term treatment (24 weeks) to achieve good clinical results; thus, other retinoids have been explored and the results of their application in pigmentary disorders have been reviewed [75]. Tarazotene, an

141

142

5 Inhibitors and Enhancers of Melanogenesis

acetylenic topical retinoid, was found to be effective towards irregular hyperpigmentation associated with photoaging [76], whereas isotretinoin did not show signiﬁcant positive results in melasma patients [272]. Topical application of β-carotene in nanothalospheres, which are special vectors able to deliver intact β-carotene into the intracellular space of melanocytes, induced good skin-lightening action with minimal side-effects [78]. Adapalene is a naphthoic acid derivative with potent retinoid activity; it controls cell proliferation and differentiation, and has signiﬁcant anti-inﬂammatory action. In a randomized clinical trial, the efﬁcacy of adapalene 0.1% gel was found to be comparable to that of tretinoin 0.05% cream in the treatment of melasma and solar lentigines, but patients using adapalene showed fewer side-effects and greater acceptability [273]. The treatment of black guinea pig skin with the combination of retinoic acid with the monobenzylether of HQ produced a complete degree of depigmentation in the majority of treated sites and reduced the average number of melanocytes [274]. A combination of the retinoid retinaldehyde 0.1% and glycolic acid 6.4% showed a higher skin-depigmenting potential than retinoic acid 0.05% in the tail skin of C57BL/6 mice [275]. 5.2.3.2.2 Chemical Peels To treat pigmentary lesions such as solar lentigines, melasma, and postinﬂammatory hyperpigmentation, superﬁcial and medium-depth peels with trichloroacetic acid (TCA), α-hydroxy acids (AHAs), such as lactic acid, glycolic acid, and salicylic acid, and tretinoin may be used mainly in fair-skinned individuals even if some clinical trials have proved their efﬁcacy also in dark-skinned patients [276]. According to the concentrations used, AHAs can induce different effects: promotion of exfoliation by decreasing corneocyte cohesion and stimulating basal layer renewal, at low concentrations; promotion of epidermolysis and melanin dispersion in the basal layer, at high concentrations. Furthermore, AHAs showed a direct inhibition of tyrosinase, without inﬂuencing mRNA or protein expression [266]. A clinical trial on TCA peels reported 40% complete regression and 50% partial regression of lentigines [277]. Lactic acid supplemented with AsA determined a general skin-whitening effect in subjects with medium to dark skin [278]. Salicylic acid is effective in inducing skin desquamation in dark-skinned individuals [79], possibly acting also by a noncompetitive inhibition of tyrosinase [279]. Owing to the low risk of inﬂammatory hyperpigmentation, glycolic acid peels can be used safely in dark-skinned individuals. A 1% tretinoin peel has been suggested as a well-tolerated and effective chemical peel for melasma as well as 70% glycolic acid [280]. According to Kligman’s opinion, however, 1% tretinoin could act as a modulator of gene expression, leading to acceleration of epidermal cell turnover, rather than as a peeling agent [281]. 5.2.3.2.3 Linoleic Acid Apart from its effect on tyrosinase degradation, linoleic acid also inﬂuences skin pigmentation by stimulating epidermal turnover and increased desquamation of melanin pigment from the epidermis. Skin-lightening capabilities of unsaturated fatty acids, linoleic acid or α-linoleic acid, have been evaluated on UV-induced

5.3 Enhancers of Melanogenesis

hyperpigmentation of brown guinea pig skin [131]. Inhibition of melanogenesis in vivo has been proposed to be related to the peroxidation of the unsaturated bonds of these molecules and to the increased epidermal turnover. 5.2.3.2.4 Licorice Extracts The compounds extracted from the roots of licorice (Glycyrrhiza glabra L.) are widely used in traditional Chinese medicine for the treatment of a number of diseases [54, 171, 282]. Some of these molecules show depigmenting properties. Among hydrophilic extracts of licorice, liquiritin, a glucoside-derived ﬂavonoid, was found to be effective for prolonged treatments of epidermic melasma [81]. The proposed mechanism of action is an acceleration of epidermis turnover and melanin dispersion induced by ﬂavone rings contained in liquiritin and its homolog isoliquiritin. Glabridin is the main component of the hydrophobic fraction of licorice, and it has been reported to inhibit UV-B-induced pigmentation and tyrosinase activity [283], although no evaluation of its efﬁcacy as a depigmenting agent has been carried out. Moreover, glabridin is able to reduce superoxide production and cyclooxygenase activity, suggesting that this compound can counteract skin inﬂammation as well as melanin biosynthesis induced by UV exposure. Licorice extracts interfere with different steps of the pigmentation process, by inhibiting tyrosinase activity in a dose-dependent manner and by removing epidermal melanin, and they have been tested for treating melasma patients with good results and very mild irritation [142].

5.3 Enhancers of Melanogenesis

Constitutive pigmentation is the result of genetic individual background that determines the phenotype of the different skin types (see Chapter 11). The facultative pigmentation is a mechanism that has evolved to protect human skin from high levels of terrestrial UV rays. Increased cutaneous eumelanin production results in skin pigmentation, providing a partial barrier to the penetration of UV and visible light. In particular, it forms a protective “nuclear cap” of melanin over the nuclei of basal keratinocytes. Eumelanin also scavenges UV-induced ROS that can damage DNA, protein, and lipids. Thus, when the melanin shield is compromised individuals are exposed to increased skin aging and cancer risk. While the photoprotective beneﬁt of tanning is quite modest, there may be noncarcinogenic pharmacological opportunities to stimulate eumelanogenesis, avoiding UV-induced genotoxic damage. To deﬁne the optimal approach for enhancing eumelanin production the signaling pathways and intrinsic/extrinsic factors that inﬂuence melanocyte proliferation or metabolism should be taken in account. After UV exposure melanin synthesis is activated via the increase of MITF, which is in turn regulated by the transcription factor SOX-9. Several signaling molecules, such as bFGF, hepatocyte growth factor (HGF), SCF, ET-1, ACTH, and α-MSH, bind their respective receptors present on melanocytes, stimulating pigment

143

144

5 Inhibitors and Enhancers of Melanogenesis

production. Other pathways involving p53, peroxisome proliferator-activated receptor (PPAR) or ERK are also involved in the melanogenic response (see Chapter 11). 5.3.1 Activation Through Receptor Mechanisms 5.3.1.1 Melanotropic Peptides MC1R is one of several proteins involved in the normal biosynthesis of melanin. α-MSH binds MC1R to stimulate both eumelanogenesis, by upregulating tyrosinase activity, and melanocyte proliferation, through activation of adenylate cyclase. Although α-MSH stimulates natural skin protection, the process requires harmful UV radiation and it is highly susceptible to proteolytic degradation. The development of a synthetic analog was therefore suggested as a suitable strategy to obtain a tanning agent, which stimulated the protective effect of eumelanogenesis without requiring UV and with a long duration of action. [Nle4-d-Phe7]-α-MSH, also called NDP-MSH or Melanotan, is one of several potent analogs of α-MSH, which has been developed by replacing the methionine at position 4 with norleucine, and the phenylalanine is racemized to the d-isomer at position 7. NDP-MSH exhibits a 10- to 100-fold increased activity respect to the endogenous α-MSH molecule, increases melanogenesis and tyrosinase activity in human melanocytes, and more speciﬁcally induces signiﬁcant increases in the eumelanin content of melanocytes while having a lesser and more varied effect on the levels of pheomelanin. Subcutaneous injection of NDP-MSH increases skin pigmentation and activates eumelanin expression, without any signiﬁcant change in pheomelanin expression, reproducing, thus, the results of in vitro studies. In humans, increased skin pigmentation occurs despite minimization of sun exposure and concomitant use of sunscreens, although it was more evident in sun-exposed areas, than in covered skin sites [284]. However this study has been performed only in subjects with skin types III or IV, avoiding the opportunity to reach any conclusion about the potential increase of eumelanin in individuals with skin types I or II, which are characterized by high levels of pheomelanin and high susceptibility to UV-induced harmful effects. In a double-blind randomized placebo-controlled study involving 65 subjects with low or high minimal erythemal dose (MED) daily subcutaneous injection of NDP-MSH for 10 days in three monthly cycles preferentially increased cutaneous melanin density in subjects with the lowest baseline skin melanin levels, and also resulted in a 50% decrease in epidermal sunburn cells and a 59% reduction in thymine dimers formation in the low MED group [285]. Considering that in humans, as in other animals, MC1R is a key point in the regulation of pigmentation phenotype and that variations of gene sequence of this protein are associated with a poor tanning response, to investigate whether the presence of alleles with diminished function would affect the binding afﬁnity and efﬁcacy of NDP-MSH represents an important topic. Earlier studies investigating key receptor residues involved in α-MSH–MC1R binding found that while the mutation of

5.3 Enhancers of Melanogenesis

conserved amino acids reduced the binding afﬁnity of α-MSH, that of NDP-MSH remained unchanged or only slightly reduced. More recently it was found that both homozygous or heterozygous cultures for several alleles, including the diminished function variants Arg142His, Arg151His, and Leu93Arg, demonstrated a signiﬁcantly reduced response to normal α-MSH and superpotent NDP-MSH compared to wild-type receptors. Fitzgerald et al. [286] investigated for the ﬁrst time the effect of NDP-MSH in human skin with a variant MC1R genotype. Their data demonstrated that NDP-MSH causes the greatest increase in skin melanin density in MC1R variant carriers (i.e., alleles associated with fair skin and red hair), suggesting that individuals with a probable increased risk of skin cancer can beneﬁt from a drug suspected to only work in wild-type MC1R individuals. Other studies have conﬁrmed that NDP-MSH treatment induces skin pigmentation both in the presence and absence of UV radiation, although UV-B radiation or sunlight appear to have a synergistic effect on the tanning response [287]. 5.3.1.2 Cytokines and Growth Factors Many studies have demonstrated that melanocyte growth and survival is regulated by keratinocyte-derived factors, such as bFGF, SCF, and ETs. In particular, SCF activates its transmembrane tyrosine kinase receptor, c-kit, promoting dimerization, autophosphorylation, and transphosphorylation of several substrates at speciﬁc tyrosine residues. After associating with the adapter protein, Grb2, an activated c-kit induces tyrosinase phosphorylation of phospholipase C-γ, which mediates the production of diacylglycerol and activates the PKC pathway. Both the SCF/c-kit and the ET-1/type B ET receptor (ETBR) pathways are crucial not only during neural crest formation and early melanocyte development, but also for melanocyte function postnatally [36]. Intradermal injection of SCF or c-kit neutralizing antibody into human xenografts was shown to regulate the number of Ki67-postive melanocytes. The synergistic effect of SCF/c-kit and ET-1/ETBR pathways in enhancing the level of DNA synthesis and activating the MAPK pathway was previously evaluated using an in vitro system, in which the combination of these two cytokines was added to human melanocyte cultures. Downregulation of SCF in vitiliginous keratinocytes was reported to be responsible for keratinocyte and melanocyte apoptosis. The treatment with either SCF or ET-1 has been shown to inﬂuence cell adhesion and melanocyte cell migration in culture, implying the plausible role of these two cytokines in regulating melanocyte function in vitiligo [36]. In addition, it has been postulated that these cytokines may inﬂuence the recruitment of melanocytes out of pigmented hair follicles in vitiligo lesions to induce repigmentation. The effect of both SCF and ET-1 in living intact human skin has been evaluated in a xenograft model using fresh Caucasian cadaveric skin on mice. Intradermal injections of the combination of these two paracrine melanogenic cytokines was shown to induce proliferation, melanogenesis, and dendritogenesis of melanocytes, suggesting that SCF administered locally in combination with ET-1 may have a potential therapeutic effect for the treatment of melanocytopenic disorders [288]. Geniposide, which is an

145

146

5 Inhibitors and Enhancers of Melanogenesis

herbal extract used in traditional Chinese medicine for the treatment of generalized vitiligo, was found to increase pigmentation via SCF/c-kit in normal human melanocytes [289]. Due to the fact that in vitiligo the presence of cytotoxic T cells targeting melanocyte antigens and an imbalance of the cytokine network were described, therapeutic approaches able to counteract immune response mediators have been investigated. In recent years, vitamin D analogs, particularly calcipotriol and tacalcitol, have been used as topical therapeutic agents in vitiligo. Vitamin D ligands are designed to target the local immune response in vitiligo, acting on speciﬁc T cell activation, mainly by inhibiting the transition of T cells from early to late G1 phase and by inhibiting the expression of several proinﬂammatory cytokines genes, such as those encoding TNF-α and interferon-γ. Vitamin D3 compounds are known to inﬂuence melanocyte maturation and differentiation, and also to upregulate melanogenesis through pathways activated by speciﬁc ligand receptors, such as ET receptor and c-kit [290]. Bone morphogenetic proteins (BMPs) are secreted signaling molecules that belong to the TGF-β superfamily. Apart from their role as stimulators of bone formation, BMPs affect other tissues, inducing various effects like proliferation that oppose differentiation and apoptosis. To date, more than 20 different BMP proteins have been identiﬁed, all sharing structural homology and interaction with speciﬁc BMP receptors. BMP effects depend on many factors including tissue concentration, BMP type, presence of antagonists, embryonic stage of the target tissue, and the type of receptors expressed by the target cells. BMP-2 has been reported to stimulate melanogenesis, without having a direct effect upon melanocyte differentiation, as demonstrated by the BMP-2- induced increase of tyrosinase mRNA, without affecting MITF, TRP-1, or TRP-2 mRNAs, indicating that BMP-2 signaling selectively targets the transcription of the tyrosinase gene [291]. Noggin is a secreted BMP antagonist that competes for binding to BMP receptors. Mice overexpressing noggin in the hair follicle epithelium have been reported to display a darker coat color than wild-type mice, demonstrating that BMP signaling inﬂuence murine melanogenesis in vivo [292]. Yaar et al. reported that normal neonatal human melanocytes and keratinocytes express the BMP receptors and produce BMP-4, suggesting that BMP antagonists can enhance skin pigmentation [293]. The extract of the human placenta, named “melagenin,” has been reported to stimulate in vitro the proliferation of melanocytes a nd the synthesis of melanin. A pilot trial evaluated the effectiveness of the topical application of the melagenin in pediatric vitiligo patients. Sixty-two out of the 366 treated subjects, ranging from 4 to 15 years old and followed up for 1 year, were characterized by depigmentation involving more than 70% of the body. The authors reported that the treatment with melagenin proved to be effective in 83% of vitiligo patients [294]. Currently, the placenta extract is used in Cuba, even if the protocol lacks external validation. The possible mechanism of action of the extract has been reported to be associated with the presence of melanocyte growth factors in placenta, including ET.

5.3 Enhancers of Melanogenesis

5.3.2 Non-Receptor-Mediated Activation 5.3.2.1 Forskolin and cAMP The skin-tanning effectiveness of α-MSH analogs is affected by the technical difﬁculties of administering peptides to humans, as well as by the requirement of a functional MC1R to activate eumelanogenesis. In direct analogy to humans, MC1R mutations in mice lead to yellow coat color and inability to tan. The major intracellular messenger used by MC1R to induce pigmentation is cAMP. Forskolin directly activates adenylate cyclase thereby increasing cAMP levels and stimulates MITF activity, reproducing the intracellular signaling pathway activated by MC1R. Due to its ability to bypass MC1R, forskolin is hypothesized to enhance pigmentation regardless of MC1R genotype. In mice expressing nonfuctional MC1R daily topical application of forskolin has been reported to stimulate the tanning response, affording a pigmentation able to counteract UV-induced damage and carcinogenesis [295]. Forskolin was also found to increase the removal of cyclobutane pyrimidine dimers and 6,4-photoproducts in human keratinocytes treated with UV-B [296]. These data conﬁrmed the ability of forskolin to reproduce the natural response of epidermis to UV rays, even in subjects with loss-of-function MC1R variants, increasing skin pigmentation and ability to repair DNA photodamage, without needing UV exposure. 5.3.2.2 Oligonucleotides and p53 Activation The small DNA fragment thymidine dinucleotide pTpT induces photoprotective responses in cultured cells and intact skin via melanogenesis, enhanced DNA repair, and induction of TNF-α. The induction and activation of the p53 tumor suppressor and transcription factor is the key signaling pathway. Larger oligonucleotides induce the pigmentation response even more efﬁciently than pTpT. The ability of these oligonucleotides to stimulate pigmentation is highly dependent on the presence of a 5′-phosphate group on the molecule [297]. 5.3.2.3 Piperin The water extract (0.1 mg/ml) of black pepper (Piper nigrum), and its alkaloid piperine, has been demonstrated to have in vitro proliferative activity, probably mediated by activation of PKC, on melanocytes. Some mouse observations have conﬁrmed the in vitro data [298]. 5.3.2.4 Lipids (Sphingolipids and Prostaglandins) The occasional report of prostaglandin E2-induced hyperpigmentation after eyedrop usage in ophthalmology suggested its possible utilization for the topical treatment of vitiligo lesions. In vitro, prostaglandin E2 has been reported to induce the expression of mRNA for bFGF and the oxidative stress-mediated glutathione depletion may decrease the prostaglandin E2 level in vitiligo epidermis. A recent trial indicates that the topical application of a gel containing prostaglandin E2 gives rise to repigmentation, mainly on the face, after 6 months of therapy. Segmental

147

148

5 Inhibitors and Enhancers of Melanogenesis

and focal vitiligo have been reported to be characterized by the highest percentage of repigmentation [299]. 5.3.2.5 Phospholipase A2 Phospholipase A2, a component of bee venom, has been considered for vitiligo repigmentation [300]. When tested in vitro, bee venom was able to promote melanocyte proliferation, dendriticity, and migration; the tyrosinase activity was also positively affected. The molecular mechanism has been supposed to be mediated by the activation of intracellular signal transduction mechanisms, in particular of PKA, ERK, and PI3K/Akt. 5.3.2.6 PPAR Activators PPARs play an important role in cellular responses. It was reported that three subtypes of PPAR are expressed in human melanocytes. The PPAR-γ agonist, ciglitazone, stimulates the melanin content of cells and cultured skin, in a tyrosinase/MITF-dependent pathway. Migration of melanocytes was increased after ciglitazone treatment [301]. 5.3.2.7 Psoralens and Photosensitizing Agents PUVA (psoralens plus UV-A) induces pigmentation that may develop without clinically evident erythema. PUVA is used for the treatment of vitiligo and for the preventive treatment of photodermatoses. In normal skin, PUVA pigmentation peaks about 7 days after exposure and may last from several weeks to months. A few PUVA exposures result in a much deeper tan than that produced by multiple exposures to solar irradiation. Psoralens belong to the furocoumarin group of compounds that are derived from the fusion of the furan ring with coumarin. These are found in a large number of plants and there are also several synthetic psoralen compounds. 8-Methoxypsoralen (8-MOP) is of plant origin, but it is also available as a synthetic drug. It is used primarily for oral and topical PUVA. The synthetic compound 4,5′,8-trimethylpsoralen (TMP, trioxsalen) is less phototoxic after oral administration, but more phototoxic when bath-water delivered. 5-MOP (bergapten) is only used in some European countries, and is less erythemogenic and not associated with gastrointestinal intolerance. Without UV radiation, psoralens intercalate in DNA double strands. After the absorption of UV-A photons the formation of 3,4- or 4′,5′-cyclobutane monoadducts with pyrimidine bases of native DNA occurs. Some psoralens, such as 8-MOP, 5-MOP, and TMP, can absorb a second photon, and this reaction leads to the formation of a bifunctional adduct that inhibits DNA replication and causes cell cycle arrest. Psoralens also stimulate melanogenesis. This involves the photoconjugation of psoralens to DNA in melanocytes followed by mitosis and subsequent proliferation of melanocytes, increased formation and melanization of melanosomes, increased transfer of melanosomes to keratinocytes, and activation and increased synthesis of tyrosinase via stimulation of cAMP activity [302].

References

References 1 Sejii, M., Shimao, K., Birbeck, M.S., and

2

3

4

5

6

7

8

9

Fitzpatrick, T.B. (1963) Subcellular localization of melanin biosynthesis. Ann. NY Acad. Sci., 100, 497–533. Kushimoto, T., Basrur, V., Valencia, J., Matsunaga, J., Vieira, W.D., Ferrans, V.J., Muller, J., Appella, E., and Hearing, V.J. (2001) A model for melanosome biogenesis based on the puriﬁcation and analysis of early melanosomes. Proc. Natl. Acad. Sci. USA, 98, 10698–10703. Raposo, G. and Marks, M.S. (2007) Melanosomes – dark organelles enlighten endosomal membrane transport. Nat. Rev. Mol. Cell Biol., 8, 786–797. Choi, W., Wolber, R., Gerwat, W., Mann, T., Batzer, J., Smuda, C., Liu, H., Kolbe, L., and Hearing, V.J. (2010) The ﬁbroblast-derived paracrine factor neuregulin-1 has a novel role in regulating the constitutive color and melanocyte function in human skin. J. Cell Sci., 123, 3102–3111. Cario-André, M., Pain, C., Gauthier, Y., Casoli, V., and Taieb, A. (2006) In vivo and in vitro evidence of dermal ﬁbroblasts inﬂuence on human epidermal pigmentation. Pigment Cell Res., 19, 434–442. Szabó, G. (1954) The number of melanocytes in human epidermis. Br. Med. J., 1, 1016–1017. Tobin, D., Quinn, A.G., Ito, S., and Thody, A.J. (1994) The presence of tyrosinase and related proteins in human epidermis and their relationship to melanin type. Pigment Cell Res., 7, 204–209. Hunt, G., Kyne, S., Ito, S., Wakamatsu, K., Todd, C., and Thody, A. (1995) Eumelanin and phaeomelanin contents of human epidermis and cultured melanocytes. Pigment Cell Res., 8, 202–208. Maeda, K., Yokokawa, Y., Hatao, M., Naganuma, M., and Tomita, Y. (1997) Comparison of the melanogenesis in human black and light brown melanocytes. J. Dermatol. Sci., 14, 199–206.

10 Szabó, G., Gerald, A.B., Pathak, M.A.,

11

12

13

14

15

16

17

18

and Fitzpatrick, T.B. (1969) Racial differences in the fate of melanosomes in human epidermis. Nature, 222, 1081–1082. Urabe, K., Nakayama, J., and Hori, Y. (1998) Mixed epidermal and dermal hypermelanoses, in The Pigmentary System: Physiology and Pathophysiology (eds J.J. Norlund, R.E. Boissy, V.J. Hearing, R.A. King, and J.P. Ortonne), Oxford University Press, New York, pp. 909–911. Virador, V., Matsunaga, N., Matsunaga, J., Valencia, J., Oldham, R.J., Kameyama, K., Peck, G.L., Ferrans, V.J., Vieira, W.D., Abdel-Malek, Z.A., and Hearing, V.J. (2001) Production of melanocyte-speciﬁc antibodies to human melanosomal proteins: expression patterns in normal human skin and in cutaneous pigmented lesions. Pigment Cell Res., 14, 289–297. Sanchez-Ferrer, A., Rodriguez-Lopez, J.N., Garcia-Canovas, F., and GarciaCarmona, F. (1995) Tyrosinase: a comprehensive review of its mechanism. Biochim. Biophys. Acta, 1247, 1–11. Halaouli, S., Asther, M., Sigoillot, J.C., Hamdi, M., and Lomascolo, A. (2006) Fungal tyrosinases: new prospects in molecular characteristics, bioengineering and biotechnological applications. J. Appl. Microbiol., 100, 219–232. Toyofuku, K., Wada, I., Valencia, J.C., Kushimoto, T.O., Ferrana, V.J., and Hearing, V.J. (2001) Oculocutaneous albinism types 1 and 3 are ER retention diseases: mutation of tyrosinase or Tyrp1 can affect the processing of both mutant and wild-type proteins. FASEB J., 15, 2149–2161. Hearing, V.J. (2000) The melanosome: the perfect model for cellular responses to the environment. Pigment Cell Res., 13, 23–34. Sarangarajan, R. and Boissy, R.E. (2001) Tyrp1 and oculocutaneous albinism type 3. Pigment Cell Res., 14, 437–444. Urabe, K., Aroca, P., Tsukamoto, K., Mascagna, D., Palumbo, A., Prota, G.,

149

150

5 Inhibitors and Enhancers of Melanogenesis

19

20

21

22

23

24

25

26

27

and Hearing, V.J. (1994) The inherent cytotoxicity of melanin precursors: a revision. Biochim. Biophys. Acta, 1221, 272–278. Hennessy, A., Oh, C., Diffey, B., Wakamatsu, K., Ito, S., and Rees, J. (2005) Eumelanin and pheomelanin concentrations in human epidermis before and after UV-B irradiation. Pigment Cell Res., 18, 220–223. Liu, Y., Hong, L., Wakamatsu, K., Ito, S., Adhyaru, B., Cheng, C.Y., Bowers, C.R., and Simon, J.D. (2005) Comparison of structural and chemical properties of black and red human hair melanosomes. Photochem. Photobiol., 81, 135–144. Branza-Nichita, N., Petrescu, A., Negroiu, G., Dwek, R., and Petrescu, S. (2000) N-Glycosylation processing and glycoprotein folding − lessons from the tyrosinase-related proteins. Chem. Rev., 100, 4697–4711. Imokawa, G. and Mishima, Y. (1984) Functional analysis of tyrosinase isozymes of cultured malignant melanoma cells during the recovery period following interrupted melanogenesis induced by glycosylation inhibitors. J. Invest. Dermatol., 83, 196–201. Halaban, R., Cheng, E., Zhang, Y., Moellmann, G., Hanlon, D., Michalak, M., Setaluri, V., and Hebert, D.N. (1997) Aberrant retention of tyrosinase in the endoplasmic reticulum mediates accelerated degradation of the enzyme and contributes to the dedifferentiated phenotype of amelanotic melanoma cells. Proc. Natl. Acad. Sci. USA, 94, 6210–6215. Hiller, M.M., Finger, A., Schweiger, M., and Wolf, D.H. (1996) ER degradation of a misfolded luminal protein by the cytosolic ubiquitin–proteasome pathway. Science, 273, 1725–1728. Hochstrasser, M. (1996) Protein degradation or regulation: Ub the judge. Cell, 22, 813–815. Hershko, A. and Ciechanover, A. (1998) The ubiquitin system. Annu. Rev. Biochem., 67, 425–479. Ando, H., Ichihashi, M., and Hearing, V.J. (2009) Role of the ubiquitin

28

29

30

31

32

33

34

35

36

37

proteasome system in regulating skin pigmentation. Int. J. Mol. Sci., 10, 4428–4434. Jackson, I.J., Budd, P., Horn, J.M., Johnson, R., Raymond, S., and Steel, K. (1994) Genetics and molecular biology of mouse pigmentation. Pigment Cell Res., 7, 73–80. García-Borrón, J.C., and Solano, F. (2002) Molecular anatomy of tyrosinase and its related proteins: beyond the histidine-bound metal catalytic center. Pigment Cell Res., 15, 162–173. Kwon, B.S. (1993) Pigmentation genes: the tyrosinase gene family and the pmel 17 gene family. J. Invest. Dermatol., 100, 134S–140S. Kwon, B.S., Haq, A.K., Pomerantz, S., and Halaban, R. (1987) Isolation and sequence of a cDNA clone for human tyrosinase that maps at the mouse c-albino locus. Proc. Natl Acad. Sci. USA, 84, 7473–7477. Shibahara, S., Tomita, Y., Tagami, H., Muller, R.M., and Cohen, T. (1988) Molecular basis for the heterogeneity of human tyrosinase. Tohoku J. Exp. Med., 156, 403–414. Riley, P.A. (2000) Tyrosinase kinetics: a semi-quantitative model of the mechanism of oxidation of monohydric and dihydric phenolic substrates. J. Theor. Biol., 203, 1–12. Olivares, C. and Solano, F. (2009) New insights into the active site structure and catalytic mechanism of tyrosinase and its related proteins. Pigment Cell Melanoma Res., 22, 750–760. Inoue, T., Shiota, Y., and Yoshizawa, K. (2008) Quantum chemical approach to the mechanism for the biological conversion of tyrosine to dopaquinone. J. Am. Chem. Soc., 130, 16890–16897. Imokawa, G. (2004) Autocrine and paracrine regulation of melanocytes in human skin and in pigmentary disorders. Pigment Cell Res., 17, 96–110. Miyamura, Y., Coelho, S.G., Wolber, R., Miller, S.A., Wakamatsu, K., Zmudzka, B.Z., Ito, S., Smuda, C., Passeron, T., Choi, W., et al. (2007) Regulation of human skin pigmentation and responses to ultraviolet radiation. Pigment Cell Res., 20, 2–13.

References 38 Yamaguchi, Y., Brenner, M., and

39

40

41

42

43

44

45

46

Hearing, V.J. (2007) The regulation of skin pigmentation. J. Biol. Chem., 282, 27557–27561. Shibahara, S., Takeda, K., Yasumoto, K., Udono, T., Watanabe, K., Saito, H., and Takahashi, K. (2001) Microphthalmiaassociated transcription factor (MITF): multiplicity in structure, function, and regulation. J. Investig. Dermatol. Symp. Proc., 6, 99–104. Cheli, Y., Ohanna, M., Ballotti, R., and Bertolotto, C. (2010) Fifteen-year quest for microphthalmia-associated transcription factor target genes. Pigment Cell Melanoma Res., 23, 27–40. Bertolotto, C., Abbe, P., Hemesath, T.J., Bille, K., Fisher, D.E., Ortonne, J.P., and Ballotti, R. (1998) Microphthalmia gene product as a signal transducer in cAMP-induced differentiation of melanocytes. J. Cell Biol., 142, 827–835. Kamaraju, A.K., Bertolotto, C., Chebath, J., and Revel, M. (2002) Pax3 downregulation and shut-off of melanogenesis in melanoma B16/F10.9 by interleukin-6 receptor signaling. J. Biol. Chem., 277, 15132–15141. Yamaguchi, Y., Morita, A., Maeda, A., and Hearing, V.J. (2009) Regulation of skin pigmentation and thickness by Dickkopf 1 (DKK1). J. Investig. Dermatol. Symp. Proc, 14, 73–75. Schepsky, A., Bruser, K., Gunnarsson, G.J., Goodall, J., Hallsson, J.H., Goding, C.R., Steingrimsson, E., and Hecht, A. (2006) The microphthalmia-associated transcription factor Mitf interacts with beta-catenin to determine target gene expression. Mol. Cell Biol., 26, 8914–8927. Levy, C., Khaled, M., and Fisher, D. (2006) MITF: master regulator of melanocyte development and melanoma oncogene. Trends Mol. Med., 12, 406–414. Lin, C., Babiarz, L., Liebel, F., Roydon Price, E., Kizoulis, M., Gendimenico, G., Fisher, D., and Seiberg, M. (2002) Modulation of microphthalmiaassociated transcription factor gene expression alters skin pigmentation. J. Invest. Dermatol., 119, 1330–1340.

47 Saha, B., Sing, S., Sarkar, C., Bera, R.,

48

49

50

51

52

53

54

55

56

Ratha, J., Tobin, D., and Bhadra, R. (2006) Activation of the Mitf promoter by lipid-stimulated activation of p38-stress signalling to CREB. Pigment Cell Res., 19, 595–605. Imokawa, G., Yada, Y., and Miyagishi, M. (1992) Endothelins secreted from human keratinocytes are intrinsic mitogens for human melanocytes. J. Biol. Chem., 267, 24675–24680. Morita, E., Lee, D.G., Sugiyama, M., and Yamamoto, S. (1994) Expression of c-kit ligand in human keratinocytes. Arch. Dermatol. Res., 286, 273–277. Halaban, R., Ghosh, S., and Baird, A. (1987) bFGF is the putative natural growth factor for human melanocytes. In Vitro Cell. Dev. Biol., 23, 47–52. Schauer, E., Trautinger, F., Köck, A., Schwarz, A., Bhardwaj, R., Simon, M., Ansel, J.C., Schwarz, T., and Luger, T.A. (1994) Proopiomelanocortin-derived peptides are synthesized and released by human keratinocytes. J. Clin. Invest., 93, 2258–2262. Kang, H.Y., Hwang, J.S., Lee, J.Y., Ahn, J.H., Kim, J.Y., Lee, E.S., and Kang, W.H. (2006) The dermal stem cell factor and c-kit are overexpressed in melasma. Br. J. Dermatol., 154, 1094–1099. Virador, V.M., Kobayashi, N., Matsunaga, J., and Hearing, V.J. (1999) A standardized protocol for assessing regulators of pigmentation. Anal. Biochem., 270, 207–219. Solano, F., Briganti, S., Picardo, M., and Ghanem, G. (2006) Hypopigmenting agents: an updated review on biological, chemical and clinical aspects. Pigment Cell Res., 90, 550–571. Yoon, T.J., Lei, T.C., Yamaguchi, Y., Batzer, J., Wolber, R., and Hearing, V.J. (2003) Reconstituted 3-dimensional human skin of various ethnic origins as an in vitro model for studies of pigmentation. Anal. Biochem., 318, 260–269. Ni-Komatsu, L., Leung, J.K., Williams, D., Min, J., Khersonsky, S.M., Chang, Y.T., and Orlow, S.J. (2005) Triazinebased tyrosinase inhibitors identiﬁed by chemical genetic screening. Pigment Cell Res., 18, 447–453.

151

152

5 Inhibitors and Enhancers of Melanogenesis 57 Ni-Komatsu, L. and Orlow, S.J. (2007)

58

59

60

61

62

63

64

65

66

Identiﬁcation of novel pigmentation modulators by chemical genetic screening. J. Invest. Dermatol., 127, 1585–1592. Maldonado, E., Hernandez, F., Lozano, C., Castro, M.E., and Navarro, R.E. (2006) The zebraﬁsh mutant vps18 as a model for vesicle-trafﬁc related hypopigmentation diseases. Pigment Cell Res., 19, 315–326. Choi, T.Y., Kim, J.H., Ko, D.H., Kim, C.H., Hwang, J.S., Ahn, S., Kim, S.Y., Kim, C.D., Lee, J.H., and Yoon, T.J. (2007) Zebraﬁsh as a new model for phenotype-based screening of melanogenic regulatory compounds. Pigment Cell Res., 20, 120–127. Gupta, A.K., Gover, M.D., Nouri, K., and Taylor, S. (2006) The treatment of melasma: a review of clinical trials. J. Am. Acad. Dermatol., 55, 1048–1065. Rajaratnam, R., Halpern, J., Salim, A., and Emmett, C. (2010) Interventions for melasma. Cochrane Database Syst. Rev., (7), CD003583. Khemis, A., Kaiafa, A., Queille-Roussel, C., Duteil, L., and Ortonne, J.P. (2007) Evaluation of efﬁcacy and safety of rucinol serum in patients with melasma: a randomized controlled trial. Br. J. Dermatol., 156, 997–1004. Huh, S.Y., Shin, J.W., Na, J.I., Huh, C.H., Youn, S.W., and Park, K.C. (2010) The efﬁcacy and safety of 4-nbutylresorcinol 0.1% cream for the treatment of melasma: a randomized controlled split-face trial. Ann. Dermatol., 22, 21–25. Boissy, R., Visscher, M., and DeLong, M.A. (2005) A novel reversible tyrosinase inhibitor with effective in vivo skin lightening potency. Exp. Dermatol., 14, 601–608. Hamed, S., Sriwiriyanont, P., deLong, M., Visscher, M., Wickett, R., and Boissy, R. (2006) Comparative efﬁcacy and safety of deoxyarbutin, a new tyrosinase-inhibiting agent. J. Cosmet. Sci., 57, 291–308. Ertam, I., Mutlu, B., Unal, I., Alper, S., Kivçak, B., and Ozer, O. (2008) Efﬁciency of ellagic acid and arbutin in melasma: a randomized, prospective,

67

68

69

70

71

72

73

74

open-label study. J. Dermatol., 35, 570–574. Jin, Y., Lee, S., Chung, M., Park, J., Park, Y.I., Cho, T., and Lee, S. (1999) Aloesin and arbutin inhibit tyrosinase activity in a synergistic manner via a different action mechanism. Arch. Pharm. Res., 22, 232–236. Battaini, G., Monzani, E., Casella, L., Santagostini, L., and Pagliarin, R. (2000) Inhibition of the catecholase activity of biomimetic dinuclear copper complexes by kojic acid. J. Biol. Inorg. Chem., 5, 262–268. Choi, Y.K., Rho, Y.K., Yoo, K.H., Lim, Y.Y., Li, K., Kim, B.J., Seo, S.J., Kim, M.N., Hong, C.K., and Kim, D.S. (2010) Effects of vitamin C vs. multivitamin on melanogenesis: comparative study in vitro and in vivo. Int. J. Dermatol., 49, 218–226. Kasraee, B., Handjani, F., Parhizgar, A., Omrani, G.R., Fallahi, M.R., Amini, M., Nikbakhsh, M., Tran, C., Hügin, A., Sorg, O., and Saurat, J.H. (2005) Topical methimazole as a new treatment for postinﬂammatory hyperpigmentation: report of the ﬁrst case. Dermatology, 211, 360–362. Kasraee, B., Safaee Ardekani, G.H., Parhizgar, A., Handjani, F., Omrani, G.R., Samani, M., Nikbakhsh, M., Tanideh, N., Eshraghian, A., Sorg, O., and Saurat, J.H. (2008) Safety of topical methimazole for the treatment of melasma. Transdermal absorption, the effect on thyroid function and cutaneous adverse effects. Skin Pharmacol. Physiol., 21, 300–305. Grimes, P.E. (2009) Management of hyperpigmentation in darker racial ethnic groups. Semin. Cutan. Med. Surg., 28, 77–85. Hakozaki, T., Minwalla, L., Zhuang, J., Chhoa, M., Matsubara, A., Miyamoto, K., Greatens, A., Hillebrand, G., Bissett, D., and Boissy, R. (2002) The effect of niacinamide on reducing cutaneous pigmentation and suppression of melanosome transfer. Br. J. Dermatol., 147, 20–31. Kang, H.Y., Valerio, L., Bahadoran, P., and Ortonne, J.P. (2009) The role of topical retinoids in the treatment of

References

75

76

77

78

79

80

81

82

83

84

85

pigmentary disorders: an evidence-based review. Am. J. Clin. Dermatol., 10, 251–260. Ortonne, J.P. (2006) Retinoid therapy of pigmentary disorders. Dermatol. Ther., 19, 28028–28028. Sefton, J., Kligman, A.M., Kopper, S.C., Lue, J.C., and Gibson, J.R. (2000) Photodamage pilot study: a double-blind, vehicle-controlled study to assess the efﬁcacy and safety of tarazotene 0.1% gel. J. Am. Acad. Dermatol., 43, 656–663. Gordon, P.R. and Gilchrest, B.A. (1989) Human melanogenesis is stimulated by diacylglycerol. J. Invest. Dermatol., 93, 700–702. Kar, H.K. (2002) Efﬁcacy of β-carotene topical application in melasma: an open clinical trial. Indian J. Dermatol. Venereol. Leprol., 68, 320–322. Grimes, P.E. (1999) The safety and efﬁcacy of salicylic acid chemical peels in darker racial-ethnic groups. Dermatol. Surg., 25, 18–22. Lee, J., Jung, K., Kim, Y.S., and Park, D. (2007) Diosgenin inhibits melanogenesis through the activation of phosphatidylinositol-3-kinase pathway (PI3K) signaling. Life Sci., 81, 249–254. Amer, M. and Metwalli, M. (2000) Topical liquiritin improves melasma. Int. J. Dermatol., 39, 299–301. Fang, D., Tsuji, Y., and Setaluri, V. (2002) Selective down-regulation of tyrosinase family gene TYRP1 by inhibition of the activity of melanocyte transcription factor, MITF. Nucleic Acids Res., 30, 3096–3106. Kim, D.S., Kim, S.Y., Chung, J.H., Kim, K.H., Eun, H.C., and Park, K.C. (2002) Delayed ERK activation by ceramide reduces melanin synthesis in human melanocytes. Cell Signal., 14, 779–785. Watabe, H., Soma, Y., Ito, M., Kawa, Y., and Mizoguchi, M. (2002) All-trans retinoic acid induces differentiation and apoptosis of murine melanocyte precursors with induction of the microphthalmia-associated transcription factor. J. Invest. Dermatol., 118, 35–42. Fernandes, S.S., Arcuri, R., MorgadoDíaz, J.A., and Benchimol, M. (2004) Increase of melanogenesis by retinoic acid: an ultrastructural and

86

87

88

89

90

91

92

93

94

morphometric study. Tissue Cell, 36, 95–105. Yoshimura, K., Tsukamoto, K., Okazaki, M., Virador, V.M., Lei, T.C., Suzuki, Y., Uchida, G., Kitano, Y., and Harii, K. (2001) Effects of all-trans retinoic acid on melanogenesis in pigmented skin equivalents and monolayer culture of melanocytes. J. Dermatol. Sci., 27 (Suppl. 1), S68–S75. Welsh, B.M., Mason, R.S., and Halliday, G.M. (1999) Topical all-trans retinoic acid augments ultraviolet radiationinduced increases in activated melanocyte numbers in mice. J. Invest. Dermatol., 112, 271–278. Ortonne, J.P. (1992) Retinoic acid and pigment cells: a review of in vitro and in vivo studies. Br. J. Dermatol., 127 (Suppl. 41), 43–47. Cario-André, M., Lepreux, S., Pain, C., Nizard, C., Noblesse, E., and Taïeb, A. (2004) Perilesional vs. lesional skin changes in senile lentigo. J. Cutan. Pathol., 31, 441–447. Sato, K., Morita, M., Ichikawa, C., Takahashi, H., and Toriyama, M. (2008) Depigmenting mechanisms of all-trans retinoic acid and retinol on B16 melanoma cells. Biosci. Biotechnol. Biochem., 72, 2589–2597. Park, H.Y., Wu, C., Yonemoto, L., Murphy-Smith, M., Wu, H., Stachur, C.M., and Gilchrest, B.A. (2006) MITF mediates cAMP-induced protein kinase C-beta expression in human melanocytes. Biochem. J., 395, 571–578. Park, H.Y., Lee, J., González, S., Middelkamp-Hup, M.A., Kapasi, S., Peterson, S., and Gilchrest, B.A. (2004) Topical application of a protein kinase C inhibitor reduces skin and hair pigmentation. J. Invest. Dermatol., 122, 159–166. Kim, D., Park, S., Kwon, S., Youn, S., and Park, K. (2004) Effects of lysophosphatidic acid on melanogenesis. Chem. Phys. Lipids, 127, 199–206. Geilen, C.C., Bektas, M., Wieder, T., and Orfanos, C.E. (1996) The vitamin D3 analogue, calcipotriol, induces sphingomyelin hydrolysis in human keratinocytes. FEBS Lett., 378, 88–92.

153

154

5 Inhibitors and Enhancers of Melanogenesis 95 Han, W.S., Yoo, J.Y., Youn, S.W., Kim,

96

97

98

99

100

101

102

D.S., Park, K.C., Kim, S.Y., and Kim, K.H. (2002) Effects of C2-ceramide on the Malme-3M melanoma cell line. J. Dermatol. Sci., 30, 10–19. Kim, D.S., Kim, S.Y., Moon, S.J., Chung, J.H., Kim, K.H., Cho, K.H., and Park, K.C. (2001) Ceramide inhibits cell proliferation through Akt/PKB inactivation and decreases melanin synthesis in Mel-Ab cells. Pigment Cell Res., 14, 110–115. Shin, Y.J., Han, C.S., Lee, C.S., Kim, H.S., Ko, S.H., Hwang, S.K., Ko, S.G., Shin, J.W., Ye, S.K., and Chung, M.H. (2010) Zeolite 4A, a synthetic silicate, suppresses melanogenesis through the degradation of microphthalmiaassociated transcription factor by extracellular signal-regulated kinase activation in B16F10 melanoma cells. Biol. Pharm. Bull., 33, 72–76. Shimoda, N., Mutou, Y., Shimura, N., Tsukimoto, M., Awaya, A., and Kojima, S. (2010) Effect of heterocyclic pyrimidine compounds on UVB-induced cell damage in human keratinocytes and on melanogenesis in mouse B16 cells. Biol. Pharm. Bull., 33, 862–868. Cohen, P. and Frame, S. (2001) The renaissance of GSK3. Nat. Rev. Mol. Cell Biol., 2, 769–776. Martínez-Esparza, M., JiménezCervantes, C., Beermann, F., Aparicio, P., Lozano, J.A., and García-Borrón, J.C. (1997) Transforming growth factor-beta1 inhibits basal melanogenesis in B16/F10 mouse melanoma cells by increasing the rate of degradation of tyrosinase and tyrosinase-related protein-1. J. Biol. Chem., 272, 3967–3972. Martínez-Esparza, M., Solano, F., and García-Borrón, J.C. (1999) Independent regulation of tyrosinase by the hypopigmenting cytokines TGF beta1 and TNF alpha and the melanogenic hormone alpha-MSH in B16 mouse melanocytes. Cell. Mol. Biol., 45, 991–1000. Kim, D.S., Park, S.H., and Park, K.C. (2004) Transforming growth factor-beta1 decreases melanin synthesis via delayed extracellular signal-regulated kinase activation. Int. J. Biochem. Cell Biol., 36, 1482–1491.

103 Swope, V.B., Abdel-Malek, Z., Kassem,

104

105

106

107

108

109

110

111

112

L.M., and Nordlund, J.J. (1991) Interleukins 1 alpha and 6 and tumor necrosis factor-alpha are paracrine inhibitors of human melanocyte proliferation and melanogenesis. J. Invest. Dermatol., 96, 180–185. Martinez-Esparza, M., JimenezCervantes, C., Solano, F., Lozano, J.A., and Garcia-Borron, J.C. (1998) Mechanisms of melanogenesis inhibition by tumor necrosis factoralpha in B16/F10 mouse melanoma cells. Eur. J. Biochem., 255, 139–146. Choi, H., Ahn, S., Lee, B.G., Chang, I., and Hwang, J.S. (2005) Inhibition of skin pigmentation by an extract of Lepidium apetalum and its possible implication in IL-6 mediated signaling. Pigment Cell Res., 18, 439–446. Vachtenheim, J. and Borovanský, J. (2010) “Transcription physiology” of pigment formation in melanocytes: central role of MITF. Exp. Dermatol., 19, 617–627. Hornyak, T.J., Jiang, S., Guzmán, E.A., Scissors, B.N., Tuchinda, C., He, H., Neville, J.D., and Strickland, F.M. (2009) Mitf dosage as a primary determinant of melanocyte survival after ultraviolet irradiation. Pigment Cell Melanoma Res., 22, 307–318. Wang, N. and Hebert, D.N. (2006) Tyrosinase maturation through the mammalian secretory pathway: bringing color to life. Pigment Cell Res., 19, 3–18. Negroiu, G., Dwek, R.A., and Petrescu, S.M. (2005) Tyrosinase-related protein-2 and -1 are trafﬁcked on distinct routes in B16 melanoma cells. Biochem. Biophys. Res. Commun., 328, 914–921. Negroiu, G., Dwek, R.A., and Petrescu, S.M. (2003) The inhibition of early N-glycan processing targets TRP-2 to degradation in B16 melanoma cells. J. Biol. Chem., 278, 27035–27042. Ando, H., Kondoh, H., Ichihashi, M., and Hearing, V.J. (2007) Approaches to identify inhibitors of melanin biosynthesis via the quality control of tyrosinase. J. Invest. Dermatol., 127, 751–761. Takahashi, H. and Parsons, P.G. (1992) Rapid and reversible inhibition of

References

113

114

115

116

117

118

119

120

tyrosinase activity by glucosidase inhibitors in human melanoma cells. J. Invest. Dermatol., 98, 481–487. Mileo, A.M., Mattei, E., Fanuele, M., Delpino, A., and Ferrini, U. (1989) Differential radiosensitivity in cultured B-16 melanoma cells following interrupted melanogenesis induced by glucosamine. Pigment Cell Res., 2, 167–170. Negroiu, G., Branza-Nichita, N., Petrescu, A.J., Dwek, R.A., and Petrescu, S.M. (1999) Protein speciﬁc Nglycosylation of tyrosinase and tyrosinase-related protein-1 in B16 mouse melanoma cells. Biochem. J., 344, 659–665. Park, J.Y., Choi, H., Hwang, J.S., Kim, J., and Chang, I.S. (2008) Enhanced depigmenting effects of N-glycosylation inhibitors delivered by pH-sensitive liposomes into HM3KO melanoma cells. J. Cosmet. Sci., 59, 139–150. Choi, H., Ahn, S., Chang, H., Cho, N., Joo, K., Lee, B., Chang, I., and Hwang, J. (2006) Inﬂuence of N-glycan processing disruption on tyrosinase and melanin synthesis in HM3KO melanoma cells. Exp. Dermatol., 16, 110–117. Terao, M., Tomita, K., Oki, T., Tabe, L., Gianni, M., and Garattini, E. (1992) Inhibition of melanogenesis by BMY-28565, a novel compound depressing tyrosinase activity in B16 melanoma cells. Biochem. Pharmacol., 43, 183–189. Franchi, J., Coutadeur, M., Marteau, C., Mersel, M., and Kupferberg, A. (2000) Depigmenting effects of calcium d-pantetheine-S-sulfonate on human melanocytes. Pigment Cell Res., 35, 165–171. Maresca, V., Flori, E., Cardinali, G., Briganti, S., Lombardi, D., Mileo, A., Paggi, M., and Picardo, M. (2006) Ferritin light chain down-modulation generates depigmentation in human metastatic melanoma cells by inﬂuencing tyrosinase maturation. J. Cell. Physiol., 206, 843–848. Imokawa, G. (1989) Analysis of initial melanogenesis including tyrosinase transfer and melanosome differentiation

121

122

123

124

125

126

127

128

129

through interrupted melanization by glutathione. J. Invest. Dermatol., 93, 100–107. Glick, B.S. and Rothman, J.E. (1987) Possible role for fatty acyl coenzyme A in intracellular protein transport. Nature, 326, 309–312. Newton, R.A., Cook, A.L., Roberts, D.W., Leonard, J.H., and Sturm, R.A. (2007) Post-transcriptional regulation of melanin biosynthetic enzymes by cAMP and resveratrol in human melanocytes. J. Invest. Dermatol., 127, 2216–2227. Ancans, J., Tobin, D.J., Hoogduijn, M.J., Smit, N.P., Wakamatsu, K., and Thody, A.J. (2010) Melanosomal pH controls rate of melanogenesis, eumelanin/ phaeomelanin ratio and melanosome maturation in melanocytes and melanoma cells. Exp. Cell Res., 268, 26–35. Watabe, H., Valencia, J.C., Yasumoto, K., Kushimoto, T., Ando, H., Muller, J., Vieira, W.D., Mizoguchi, M., Appella, E., and Hearing, V.J. (2004) Regulation of tyrosinase processing and trafﬁcking by organellar pH and by proteasome activity. J. Biol. Chem., 279, 7971–7981. Ni-Komatsu, L., Tong, C., Chen, G., Brindzei, N., and Orlow, S.J. (2008) Identiﬁcation of quinolines that inhibit melanogenesis by altering tyrosinase family trafﬁcking. Mol. Pharmacol., 74, 1576–1586. Dahlmann, B., Becher, B., Sobek, A., Ehlers, C., Kopp, F., and Kuehn, L. (1993) In vitro activation of the 20S proteasome. Enzyme Protein, 47, 274–284. Rivett, A.J. (1993) Proteasomes: multicatalytic proteinase complexes. Biochem. J., 291, 1–10. Ando, H., Watabe, H., Valencia, J.C., Yasumoto, K., Furumura, M., Funasaka, Y., Oka, M., Ichihashi, M., and Hearing, V.J. (2004) Fatty acids regulate pigmentation via proteasomal degradation of tyrosinase: a new aspect of ubiquitin–proteasome function. J. Biol. Chem., 279, 15427–15433. Ando, H., Funasaka, Y., Oka, M., Ohashi, A., Furumura, M., Matsunaga, J., Matsunaga, N., Hearing, V.J., and Ichihashi, M. (1999) Possible

155

156

5 Inhibitors and Enhancers of Melanogenesis

130

131

132

133

134

135

136

137

138

involvement of proteolytic degradation of tyrosinase in the regulatory effect of fatty acids on melanogenesis. J. Lipid Res., 40, 1312–1316. Ando, H., Itoh, A., Mishima, Y., and Ichihashi, M. (1995) Correlation between the number of melanosomes, tyrosinase mRNA levels, and tyrosinase activity in cultured murine melanoma cells in response to various melanogenesis regulatory agents. J. Cell Physiol., 163, 608–614. Ando, H., Ryu, A., Hashimoto, A., Oka, M., and Ichihashi, M. (1998) Linoleic acid and alpha-linolenic acid lightens ultraviolet-induced hyperpigmentation of the skin. Arch. Dermatol. Res., 290, 375–381. Kageyama, A., Oka, M., Okada, T., Nakamura, S., Ueyama, T., Saito, N., Hearing, V.J., Ichihashi, M., and Nishigori, C. (2004) Downregulation of melanogenesis by phospholipase D2 through the ubiquitin proteasomemediated degradation of tyrosinase. J. Biol. Chem., 279, 27774–27780. Maeda, K., Tomita, Y., Naganuma, M., and Tagami, H. (1996) Phospholipases induce melanogenesis in organ-cultured skin. Photochem. Photobiol., 64, 220–223. Menter, J.M., Etemadi, A.A., Chapman, W., Hollins, T.D., and Willis, I. (1993) In vivo depigmentation by hydroxybenzene derivatives. Melanoma Res., 3, 443–449. Passi, S. and Nazzaro-Porro, M. (1981) Molecular basis of substrate and inhibitory speciﬁcity of tyrosinase: phenolic compounds. Br. J. Dermatol., 104, 659–665. Palumbo, A., d’Ischia, M., Misuraca, G., and Prota, G. (1991) Mechanism of inhibition of melanogenesis by hydroquinone. Biochim. Biophys. Acta, 1073, 85–90. Verallo-Rowell, V.M., Verallo, V., Graupe, K., Lopez-Villafuerte, L., and Garcia-Lopez, M. (1989) Double-blind comparison of azelaic acid and hydroquinone in the treatment of melasma. Acta Derm. Venereol. Suppl., 143, 58–61. Yang, F. and Boissy, R.E. (1999) Effects of 4-tertiary butylphenol on the

139

140

141

142

143

144

145

146

147

148

149

tyrosinase activity in human melanocytes. Pigment Cell Res., 12, 237–245. Penney, K.B., Smith, C.J., and Allen, J.C. (1984) Depigmenting action of hydroquinone depends on disruption of fundamental cell processes. J. Invest. Dermatol., 82, 308–310. Briganti, S., Camera, E., and Picardo, M. (2003) Chemical and instrumental approaches to treat hyperpigmentation. Pigment Cell Res., 16, 101–110. Draelos, Z. (2007) Skin lightening preparations and the hydroquinone controversy. Dermatol. Ther., 20, 308–313. Picardo, M. and Carrera, M. (2007) New and experimental treatments of cloasma and other hypermelanoses. Dermatol. Clin., 25, 353–362. Ennes, S.B.P., Paschoalick, R.C., and Mota de Avelar, A.M. (2000) A double-blind, comparative, placebocontrolled study of the efﬁcacy and tolerability of 4% hydroquinone as a depigmenting agent in melasma. J. Dermatol. Treat., 11, 173–179. Kamau, P. and Jordan, R.B. (2002) Kinetic study of the oxidation of catechol by aqueous copper (II). Inorg. Chem., 41, 3076–3083. Kim, Y.J., Woo, H.D., Kim, B.M., Lee, Y.J., Kang, S.J., Cho, Y.H., and Chung, H.W. (2009) Risk assessment of hydroquinone: differential responses of cell growth and lethality correlated to hydroquinone concentration. J. Toxicol. Environ. Health A, 72, 1272–1278. Ortonne, J.P. and Passeron, T. (2005) Melanin pigmentary disorders: treatment update. Dermatol. Clin., 23, 209–226. Bolognia, J.L., Sodi, S.A., Osber, M.P., and Pawelek, J.M. (1995) Enhancement of the depigmenting effect of hydroquinone by cystamine and buthionine sulfoximine. Br. J. Dermatol., 133, 349–357. Guevara, I.L. and Pandya, A.G. (2001) Melasma treated with hydroquinone, tretinoin, and a ﬂuorinated steroid. Int. J. Dermatol., 40, 212–215. Gaskell, M., McLuckie, K.I., and Farmer, P.B. (2005) Genotoxicity of the benzene

References

150

151

152

153

154

155

156

157

158

metabolites para-benzoquinone and hydroquinone. Chem. Biol. Interact., 153–154, 267–270. Fenoll, L.G., Rodríguez-López, J.N., Varón, R., García-Ruiz, P.A., GarcíaCánovas, F., and Tudela, J. (2000) Action mechanism of tyrosinase on meta- and para-hydroxylated monophenols. Biol. Chem., 381, 313–320. Thörneby-Andersson, K., Sterner, O., and Hansson, C. (2000) Tyrosinasemediated formation of a reactive quinone from the depigmenting agents, 4-tert-butylphenol and 4-tertbutylcatechol. Pigment Cell Res., 13, 33–38. Yang, F., Sarangarajan, R., Le Poole, I.C., Medrano, E.E., and Boissy, R.E. (2000) The cytotoxicity and apoptosis induced by 4-tertiary butylphenol in human melanocytes are independent of tyrosinase activity. J. Invest. Dermatol., 114, 157–164. Jimbow, K. (1991) N-Acetyl-4-Scysteaminylphenol as a new type of depigmenting agent for the melanoderma of patients with melasma. Arch. Dermatol., 127, 1528–1534. Ferguson, J., Rogers, P.M., Kelland, L.R., and Robins, D.J. (2005) Synthesis and antimelanoma activity of sterically congested tertiary amide analogues of N-acetyl-4-S-cysteaminylphenol. Oncol. Res., 15, 87– 94. Moridani, M.Y. (2006) Biochemical basis of 4-hydroxyanisole induced cell toxicity towards B16-F0 melanoma cells. Cancer Lett., 243, 235–245. Espín, J.C., Varón, R., Tudela, J., and García-Cánovas, F. (1997) Kinetic study of the oxidation of 4-hydroxyanisole catalyzed by tyrosinase. Biochem. Mol. Biol. Int., 41, 1265–1276. Rodriguez-Vicente, J., Vicente-Ortega, V., Canteras-Jordana, M., and CalderonRubiales, F. (1997) Relationship between 4-hydroxyanisole toxicity and dopa oxidase activity for three melanoma cell lines. Melanoma Res., 7, 373–381. Fleischer, A.B., Jr, Schwartzel, E.H., Colby, S.I., and Altman, D.J. (2000) The combination of 2% 4-hydroxyanisole (Mequinol) and 0.01% tretinoin is effective in improving the appearance of

159

160

161

162

163

164

165

166

solar lentigines and related hyperpigmented lesions in two double-blind multicenter clinical studies. J. Am. Acad. Dermatol., 42, 459–467. Jimbow, M., Marusyk, H., and Jimbow, K. (1995) The in vivo melanocytotoxicity and depigmenting potency of N-2,4acetoxyphenyl thioethyl acetamide in the skin and hair. Br. J. Dermatol., 133, 526–536. Gili, A., Thomas, P.D., Ota, M., and Jimbow, K. (2000) Comparison of in vitro cytotoxicity of N-acetyl and N-propionyl derivatives of phenolic thioether amines in melanoma and neuroblastoma cells and the relationship to tyrosinase and tyrosine hydroxylase enzyme activity. Melanoma Res., 10, 9–15. Njoo, M.D. and Westerhof, W. (2001) Vitiligo. Pathogenesis and treatment. Am. J. Clin.Dermatol., 2, 167–181. Fukuda, Y., Nagano, M., Tsukamoto, K., and Futatsuka, M. (1998) In vitro studies on the depigmenting activity of 4-(p-hydroxyphenyl)-2-butanone. J. Occup. Health, 40, 137–142. Kim, D.S., Kim, S.Y., Park, S.H., Choi, Y.G., Kwon, S.B., Kim, M.K., Na, J.I., Youn, S.W., and Park, K.C. (2005) Inhibitory effects of 4-n-butylresorcinol on tyrosinase activity and melanin synthesis. Biol. Pharm. Bull., 28, 2216–2219. Kim, Y.J., No, J.K., Lee, J.H., and Chung, H.Y. (2006) 3,4-dihydroxyacetophenone: inhibition of tyrosinase and MITF. Biosci. Biotechnol. Biochem., 70, 532–534. Huh, S.Y., Shin, J.W., Na, J.I., Huh, C.H., Youn, S.W., and Park, K.C. (2010) Efﬁcacy and safety of liposomeencapsulated 4-n-butylresorcinol 0.1% cream for the treatment of melasma: a randomized controlled split-face trial. J. Dermatol., 37, 311–315. Kurosu, J., Sato, T., Yoshida, K., Tsugane, T., Shimura, S., Kirimura, K., Kino, K., and Usami, S. (2002) Enzymatic synthesis of alpha-arbutin by alpha-anomer-selective glucosylation of hydroquinone using lyophilized cells of Xanthomonas campestris WU-9701. J. Biosci. Bioeng., 93, 328–330.

157

158

5 Inhibitors and Enhancers of Melanogenesis 167 Sugimoto, K., Nomura, K., Nishimura,

168

169

170

171

172

173

174

175

T., Kiso, T., Sugimoto, K., and Kuriki, T. (2005) Syntheses of alpha-arbutin-alphaglycosides and their inhibitory effects on human tyrosinase. J. Biosci. Bioeng., 99, 272–276. Nakajima, M., Shinoda, I., Fukuwatari, Y., and Hayasawa, H. (1998) Arbutin increases the pigmentation of cultured human melanocytes through mechanisms other than the induction of tyrosinase activity. Pigment Cell Res., 11, 12–17. Funayama, M., Arakawa, H., Yamamoto, R., Nishino, T., Shin, T., and Murao, S. (1995) Effects of alpha- and beta-arbutin on activity of tyrosinases from mushroom and mouse melanoma. Biosci. Biotechnol. Biochem., 59, 143–144. Sugimoto, K., Nishimura, T., Nomura, K., Sugimoto, K., and Kuriki, T. (2004) Inhibitory effects of alpha-arbutin on melanin synthesis in cultured human melanoma cells and a three-dimensional human skin model. Biol. Pharm. Bull., 27, 510–514. Parvez, S., Kang, M., Chung, H.-S., Cho, C., Hong, M.-C., Shin, M.-K., and Bae, H. (2006) Survey and mechanism of skin depigmentation and lightening agents. Phytother. Res., 20, 921–934. Chakraborty, A., Funasaka, Y., Komoto, M., and Ichihashi, M. (1998) Effect of arbutin on melanogenic proteins in human melanocytes. Pigment Cell Res., 11, 206–212. Curto, E.V., Kwong, C., Hermersdörfer, H., Glatt, H., Santis, C., Virador, V., Hearing, V.J., Jr, and Dooley, T.P. (1999) Inhibitors of mammalian melanocyte tyrosinase: in vitro comparisons of alkyl esters of gentisic acid with other putative inhibitors. Biochem. Pharmacol., 57, 663–672. Sugimoto, K., Nishimura, T., Nomura, K., Sugimoto, K., and Kuriki, T. (2003) Syntheses of arbutin-alpha-glycosides and a comparison of their inhibitory effects with those of alpha-arbutin and arbutin on human tyrosinase. Chem. Pharm. Bull., 51, 798–801. Chawla, S., deLong, M., Visscher, M., Wickett, R., Manga, P., and Boissy, R. (2008) Mechanism of tyrosinase

176

177

178

179

180

181

182

183

184

185

186

inhibition by deoxyArbutin and its second-generation derivatives. Br. J. Dermatol., 159, 1267–1274. Ebanks, J.P., Wickett, R.R., and Boissy, R.E. (2009) Mechanisms regulating skin pigmentation: the rise and fall of complexion coloration. Int. J. Mol. Sci., 10, 4066–4087. Nakagawa, M. and Kawai, K. (1995) Contact allergy to kojic acid in skin care products. Contact Dermatitis, 32, 9–13. Takizawa, T., Mitsumori, K., Tamura, T., Nasu, M., Ueda, M., Imai, T., and Hirose, M. (2003) Hepatocellular tumor induction in heterozygous p53-deﬁcient CBA mice by a 26-week dietary administration of kojic acid. Toxicol. Sci., 73, 287–293. Chang, T.S. (2009) An updated review of tyrosinase inhibitors. Int. J. Mol. Sci., 10, 2440–2475. Moon, K.Y., Ahn, K.S., Lee, J., and Kim, Y.S. (2001) Kojic acid, a potential inhibitor of NF-kappaB activation in transfectant human HaCaT and SCC-13 cells. Arch. Pharm. Res., 24, 307–311. Draelos, Z.D., Yatskayer, M., Bhushan, P., Pillai, S., and Oresajo, C. (2010) Evaluation of a kojic acid, emblica extract, and glycolic acid formulation compared with hydroquinone 4% for skin lightening. Cutis, 86, 153–158. Rho, H.S., Ahn, S.M., Yoo, D.S., Kim, M.K., Cho, D.H., and Cho, J.Y. (2010) Kojyl thioether derivatives having both tyrosinase inhibitory and antiinﬂammatory properties. Bioorg. Med. Chem. Lett., 20, 6569–6571. Noh, J.M., Kwak, S.Y., Seo, H.S., Seo, J.H., Kim, B.G., and Lee, Y.S. (2009) Kojic acid–amino acid conjugates as tyrosinase inhibitors. Bioorg. Med. Chem. Lett., 19, 5586–5589. Noh, J.M., Kwak, S.Y., Kim, D.H., and Lee, Y.S. (2007) Kojic acid–tripeptide amide as a new tyrosinase inhibitor. Biopolymers, 88, 300–307. Lee, Y.S., Park, J.H., Kim, M.H., Seo, S.H., and Kim, H.J. (2006) Synthesis of tyrosinase inhibitory kojic acid derivative. Arch. Pharm. Chem. Life Sci., 339, 111–114. Kim, D.H., Hwang, J.S., Baek, H.S., Kim, K.J., Lee, B.G., Chang, I., Kang,

References

187

188

189

190

191

192

193

194

195

196

H.H., and Lee, O.S. (2003) 5-[(3-Aminopropyl)phosphinooxy]-2(hydroxymethyl)-4H-1-pyran-4-on as a novel whitening agent. Chem. Pharm. Bull. (Tokyo), 51, 113–116. Dooley, T.P., Gadwood, R.C., Kilgore, K., and Thomasco, L.M. (1994) Development of an in vitro primary screen for skin depigmentation and antimelanoma agents. Skin Pharmacol., 7, 188–200. Kim, Y.J. and Uyama, H. (2005) Tyrosinase inhibitors from natural and synthetic sources: structure, inhibition mechanism and perspective for the future. Cell. Mol. Life Sci., 62, 1707–1723. Parvez, S., Kang, M., Chung, H., and Bae, H. (2007) Naturally occuring tyrosinase inhibitors: mechanism and application in skin health, cosmetics and agriculture industries. Phytother. Res., 21, 805–816. Nazzaro-Porro, M., Passi, S., Zina, G., and Breathnach, A.S. (1990) The depigmenting effect of azelaic acid. Arch Dermatol., 126, 1649–1651. Passi, S., Picardo, M., Mingrone, G., Breathnach, A.S., and Nazzaro-Porro, M. (1989) Azelaic acid – biochemistry and metabolism. Acta Derm. Venereol. Suppl., 143, 8–13. Nazzaro-Porro, M. (1987) Azelaic acid. J. Am. Acad. Dermatol., 17, 1033–1041. Breathnach, A.S. (1996) Melanin hyperpigmentation of skin: melasma, topical treatment with azelaic acid, and other therapies. Cutis, 57, 36–45. Hermanns, J.F., Petit, L., PiérardFranchimont, C., Paquet, P., and Piérard, G.E. (2002) Assessment of topical hypopigmenting agents on solar lentigines of Asian women. Dermatology, 204, 281–286. Nazzaro-Porro, M., Passi, S., Zina, G., and Breathnach, A.S. (1989) Ten year’s experience of treating lentigo maligna with topical azelaic acid. Acta Derm. Venereol. Suppl., 143, 49–57. Sarkar, R., Bhalla, M., and Kanwar, A.J. (2002) A comparative study of 20% azelaic acid cream monotherapy versus a sequential therapy in the treatment of

197

198

199

200

201

202

203

204

205

206

melasma in dark-skinned patients. Dermatology, 205, 249–254. Fitton, A. and Goa, K.L. (1991) Azelaic acid. A review of its pharmacological properties and therapeutic efﬁcacy in acne and hyperpigmentary skin disorders. Drugs, 41, 780–798. Yu, J., Wang, L., Walzem, R.L., Miller, E.G., Pike, L.M., and Patil, B.S. (2005) Antioxidant activity of citrus limonoids, ﬂavonoids and coumarins. J. Agric. Food Chem., 53, 2009–2014. Kubo, I., Kinst-Hori, I., Kubo, Y., Yamagiwa, Y., Kamikawa, T., and Haraguchi, H. (2000) Molecular design of antibrowning agents. J. Agric. Food Chem., 48, 1393–1399. Jones, K., Hughes, J., Hong, M., Jia, Q., and Orndorff, S. (2002) Modulation of melanogenesis by aloesin: a competitive inhibitor of tyrosinase. Pigment Cell Res., 15, 335–340. Zhu, W. and Gao, J. (2008) The use of botanical extracts as topical skinlightening agents for the improvement of skin pigmentation disorders. J. Investig. Dermatol. Symp. Proc., 13, 20–24. Wang, Z., Li, X., Yang, Z., He, X., Tu, J., and Zhang, T. (2008) Effects of aloesin on melanogenesis in pigmented skin equivalents. Int. J. Cosmet. Sci., 30, 121–130. Shimogaki, H., Tanaka, Y., Tamai, H., and Masuda, M. (2000) In vitro and in vivo evaluation of ellagic acid on melanogenesis inhibition. Int. J. Cosmet. Sci., 22, 291–303. Yoshimura, M., Watanabe, Y., Kasai, K., Yamakoshi, J., and Koga, T. (2005) Inhibitory effect of an ellagic acid-rich pomegranate extract on tyrosinase activity and ultraviolet-induced pigmentation. Biosci. Biotechnol. Biochem., 69, 2368–2373. Kasai, K., Yoshimura, M., Koga, T., Arii, M., and Kawasaki, S. (2006) Effects of oral administration of ellagic acid-rich pomegranate extract on ultravioletinduced pigmentation in the human skin. J. Nutr. Sci. Vitaminol., 52, 383–388. Kim, Y.M., Yun, J., Lee, C.K., Lee, H., Min, K.R., and Kim, Y. (2002)

159

160

5 Inhibitors and Enhancers of Melanogenesis

207

208

209

210

211

212

213

214

215

Oxyresveratrol and hydroxystilbene compounds. J. Biol. Chem., 277, 16340–16344. Ohguchi, K., Tanaka, T., Ilyya, I., Ito, T., Iinuma, M., Matsumoto, K., Asao, Y., and Nozawa, Y. (2003) Gnetol as a potent tyrosinase inhibitor from genus Gnetum. Biosci. Biotechnol. Biochem., 67, 663–665. Ohguchi, K., Tanaka, T., Kido, T., Baba, K., Iinuma, M., Matsumoto, K., Akao, Y., and Nozawa, Y. (2000) Effects of hydroxystilbene derivatives on tyrosinase activity. Biochem. Biophys. Res. Commun., 307, 861–863. Lee, K.T., Lee, K.S., Jeong, J.H., Jo, B.K., Heo, M.Y., and Kim, H.P. (2003) Inhibitory effects of Ramulus mori extracts on melanogenesis. J. Cosmet. Sci., 54, 133–142. Barber, J., Townsend, D., David, P., Olds, M.S., and King, R.A. (1984) Dopachrome oxidoreductase: a new enzyme in the pigment pathway. J. Invest. Dermatol., 83, 145–149. Fang, J., Han, Q., Johnson, J.K., Christensen, B.M., and Li, J. (2002) Functional expression and characterization of Aceds aegypi dopachrome conversion enzyme. Biochem. Biophys. Res. Commun., 290, 287–293. Choi, S.Y., Kim, S., Hwang, J.S., Lee, B.G., Kim, H., and Kim, S.Y. (2004) Benzylamide derivative compound attenuates the ultraviolet B-induced hyperpigmentation in the brownish guinea pig skin. Biochem. Pharmacol., 67, 707–715. Choi, S.Y., Hwang, J.S., Kim, S., and Kim, S.Y. (2006) Synthesis, discovery and mechanism of 2,6-dimethoxy-N-(4methoxyphenyl)benzamide as potent depigmenting agent in the skin. Biochem. Biophys. Res. Commun., 349, 39–49. Sato, K. and Toriyama, M. (2009) Effect of pyrroloquinoline quinone (PQQ) on melanogenic protein expression in murine B16 melanoma. J. Dermatol. Sci., 53, 140–145. Karg, E., Odh, G., Wittbjer, A., Rosengren, E., and Rorsman, H. (1993) Hydrogen peroxide as inducer of

216

217

218

219

220

221

222

223

elevated tyrosinase level in melanoma cells. J. Invest. Dermatol., 100, 209s–213s. Elmore, A.R. (2005) Final report of the safety assessment of l-ascorbic acid, calcium ascorbate, magnesium ascorbate, magnesium ascorbyl phosphate, sodium ascorbate, and sodium ascorbyl phosphate as used in cosmetics. Int. J. Toxicol., 24, 51–111. Kameyama, K., Sakai, C., Kondoh, S., Yonemoto, K., Nishiyama, S., Tagawa, M., Murata, T., Ohnuma, T., Quigley, J., Dorsky, A., Bucks, D., and Blanock, K. (1996) Inhibitory effect of magnesium l-ascorbyl-2-phosphate on melanogenesis in vitro and in vivo. J. Am. Acad. Dermatol., 34, 29–33. Kobayashi, S., Takehana, M., and Itoh, S. (1996) Protective effect of magnesium-l-ascorbyl-2 phosphate against skin damage induced by UV-B irradiation. Photochem. Photobiol., 64, 224–228. Matsuda, S., Shibayama, H., Hisama, M., Ohtsuki, M., and Iwaki, M. (2008) Inhibitory effects of a novel ascorbic derivative, disodium isostearyl 2-O-lascorbyl phosphate on melanogenesis. Chem. Pharm. Bull., 56, 292–297. Funasaka, Y., Chakraborty, A.K., Komoto, M., Ohashi, A., and Ichihashi, M. (1999) The depigmenting effect of α-tocopheryl ferulate on human melanoma cells. Br. J. Dermatol., 141, 20–29. Funasaka, Y., Komoto, M., and Ichihashi, M. (2000) Depigmenting effect of α-tocopheryl ferulate on normal melanocytes. Pigment Cell Res., 13 (Suppl. 8), 170–174. Shimizu, K., Kondo, R., Sakai, K., Takeda, N., Nagahata, T., and Oniki, T. (2001) Novel vitamin E derivative with 4-substituted resorcinol moiety has both antioxidant and tyrosinase inhibitory properties. Lipids, 36, 1321–1326. Nishiyama, T., Ohnishi, J., and Hashiguchi, Y. (2001) Fused heterocyclic antioxidants: antioxidative activities of hydrocumarins in a homogeneous solution. Biosci. Biotechnol. Biochem., 65, 1127–1133.

References 224 Yamamura, T., Onishi, J., and

225

226

227

228

229

230

231

232

233

234

Nishiyama, T. (2002) Antimelanogenic activity of hydrocumarins in cultured normal human melanocytes by stimulating intracellular glutathione synthesis. Arch. Dermatol. Res., 294, 349–354. Fujimori, H., Hisama, M., Shibayama, H., Kawase, A., and Iwaki, M. (2010) Inhibitory effects of phytoncide solution on melanin biosynthesis. Biosci. Biotechnol. Biochem., 74, 918–922. Yokozawa, T. and Kim, Y.J. (2007) Piceatannol inhibits melanogenesis by its antioxidative actions. Biol. Pharm. Bull., 30, 2007–2011. Kim, Y.J., Kang, K.S., and Yokozawa, T. (2008) The anti-melanogenic effect of pycnogenol by its anti-oxidative actions. Food Chem. Toxicol., 46, 2466–2471. Kim, Y.J. (2007) Antimelanogenic and antioxidant properties of gallic acid. Biol. Pharm. Bull., 30, 1052–1055. Roy, S. and Packer, L. (1998) Redox regulation of cell functions by alphalipoate: biochemical and molecular aspects. Biofactors, 8, 17–21. Podda, M., Zollner, T.M., GrundmannKollmann, M., Thiele, J.J., Packer, L., and Kaufmann, R. (2001) Activity of alpha-lipoic acid in the protection against oxidative stress in skin. Curr. Probl. Dermatol., 29, 43–51. Saliou, C., Kitazawa, M., McLaughlin, L., Yang, J.P., Lodge, J.K., Tetsuka, T., Iwasaki, K., Cillard, J., Okamoto, T., and Packer, L. (1999) Antioxidants modulate acute solar ultraviolet radiation-induced NF-kappa-B activation in a human keratinocyte cell line. Free Radic. Biol. Med., 26, 174–183. Kim, J.H., Sim, G.S., Bae, J.T., Oh, J.Y., Lee, G.S., Lee, D.H., Lee, B.C., and Pyo, H.B. (2008) Synthesis and antimelanogenic effects of lipoic acidpolyethylene glycol ester. J. Pharm. Pharmacol., 60, 863–870. Ito, S. and Prota, G. (1977) A facile one-step synthesis of cysteinyldopas using mushroom tyrosinase. Experentia, 33, 118–119. Benedetto, J.P., Ortonne, J.P., Voulot, C., Khatchadourian, C., Prota, G., and Thivolet, J. (1982) Role of thiol

235

236

237

238

239

240

241

242

243

compounds in mammalian melanin pigmentation. II. Glutathione and related enzymatic activities. J. Invest. Dermatol., 79, 422–424. Qiu, L., Zhang, M., Sturm, R.A., Gardiner, B., Tonks, I., Kay, G., and Parsons, P.G. (2000) Inhibition of melanin synthesis by cystamine in human melanoma cells. J. Invest. Dermatol., 114, 21–27. Hwang, J.S., Choi, H., Rho, H.S., Shin, H.J., Kim, D.H., Lee, J., Lee, B.G., and Chang, I. (2004) Pigment-lightening effect of N,N′-dilinoleylcystamine on human melanoma cells. Br. J. Dermatol., 150, 39–46. Okun, M.R. (1967) Peroxidase activity in normal and neoplastic melanocytes. J. Invest. Dermatol., 48, 461–465. Okun, M.R., Edelstein, L.M., Or, N., Hamada, G., and Donnellan, B. (1970) The role of peroxidase vs. the role of tyrosinase in enzymatic conversion of tyrosine to melanin in melanocytes, mast cells and eosinophils. J. Invest. Dermatol., 55, 1–12. d’Ischia, M., Napolitano, A., and Prota, G. (1991) Peroxidase as an alternative to tyrosinase in the oxidative polymerization of 5,6-dihydroxyindoles to melanin(s). Biochim. Biophys. Acta, 1073, 423–430. Nappi, A.J. and Vass, E. (1996) Hydrogen peroxide generation associated with the oxidations of the eumelanin precursors 5,6-dihydroxyindole and 5,6-dihydroxyindole-2-carboxylic acid. Melanoma Res., 6, 341–349. Chaubal, V.A., Nair, S.S., Ito, S., Wakamatsu, K., and Mojamdar, M.V. (2002) Gamma-glutamyl transpeptidase and its role in melanogenesis: redox reactions and regulation of tyrosinase. Pigment Cell Res., 15, 420–425. Schallreuter, K.U. and Wood, J.M. (1989) Free radical reduction in the human epidermis. Free Radic. Biol. Med., 6, 519–532. Lo, Y.Y., Wong, J.M., and Cruz, T.F. (1996) Reactive oxygen species mediate cytokine activation of c-Jun NH2terminal kinases. J. Biol. Chem., 271, 15703–15707.

161

162

5 Inhibitors and Enhancers of Melanogenesis 244 Meier, B., Radeke, H.H., Selle, S.,

245

246

247

248

249

250

251

252

Younes, M., Sies, H., Resch, K., and Habermehl, G.G. (1989) Human ﬁbroblasts release reactive oxygen species in response to interleukin-1 or tumour necrosis factor-alpha. Biochem. J., 263, 539–545. Thannickal, V.J. and Fanburg, B.L. (1995) Activation of an H2O2-generating NADH oxidase in human lung ﬁbroblasts by transforming growth factor beta 1. J. Biol. Chem., 270, 30334–30338. Jiménez-Cervantes, C., MartínezEsparza, M., Pérez, C., Daum, N., Solano, F., and García-Borrón, J.C. (2001) Inhibition of melanogenesis in response to oxidative stress: transient downregulation of melanocyte differentiation markers and possible involvement of microphthalmia transcription factor. J. Cell Sci., 114, 2335–2344. Mastore, M., Kohler, L., and Nappi, A.J. (2005) Production and utilization of hydrogen peroxide associated with melanogenesis and tyrosinase-mediated oxidations of DOPA and dopamine. FEBS J., 272, 2407–2415. Kasraee, B. (2002) Peroxidase-mediated mechanisms are involved in the melanocytotoxic and melanogenesisinhibiting effects of chemical agents. Dermatology, 205, 329–339. Kasraee, B. (2002) Depigmentation of brown Guinea pig skin by topical application of thimazole. J. Invest. Dermatol., 118, 205–207. Minwalla, L., Zhao, Y., Cornelius, J., Babcock, G., Wickett, R., Le Poole, I., and Boissy, R. (2001) Inhibition of melanosome transfer from melanocytes to keratinocytes by lectins and neoglycoproteins in an in vitro model system. Pigment Cell Res., 14, 185–194. Ito, Y., Kanamaru, A., and Tada, A. (2006) Centaureidin promotes dendrite retraction of melanocytes by activating Rho. Biochim. Biophys. Acta, 1760, 487–494. Ito, Y., Kanamaru, A., and Tada, A. (2006) Effects of methylophiopogonanone B on

253

254

255

256

257

258

259

260

261

melanosome transfer and dendrite retraction. J. Dermatol. Sci., 42, 68–70. Lin, J., Chiang, H., Lin, Y., and Wen, K. (2008) Natural products with skinwhitening effects. J. Food Drug Anal., 16, 1–10. Greatens, A., Hakozaki, T., Koshoffer, A., Epstein, H., Schwemberger, S., Babcock, G., Bissett, D., Takiwaki, H., Arase, S., Wickett, R., and Boissy, R. (2005) Effective inhibition of melanosome transfer to keratinocytes by lectins and niacinamide is reversible. Exp. Dermatol., 14, 498–508. Bissett, D., Oblong, J., and Berge, C. (2005) Niacinamide: a B vitamin that improves aging facial skin appearance. Dermatol. Surg., 31, 860–865. Bissett, D., Miyamoto, K., Sun, P., Li, J., and Berge, C. (2004) Topical niacinamide reduces yellowing, wrinkling, red blotchiness, and hyperpigmented spots in aging facial skin. Int. J. Cosmet. Sci., 26, 231–238. Bissett, D., Oblong, J., Saud, A., Berge, C., Trejo, A., and Biedermann, K. (2003) Topical niacinamide provides skin aging appearance beneﬁts while enhancing barrier function. J. Clin. Dermatol., 32S, 9–18. Van Den Bossche, K., Naeyaert, J., and Lambert, J. (2006) The quest for the mechanism of melanin transfer. Trafﬁc, 7, 769–778. Seiberg, M., Paine, C., Sharlow, E., Andrade-Gordon, P., Costanzo, M., Eisinger, M., and Shapiro, S. (2000) The protease-activated receptor 2 regulates pigmentation via keratinocyte– melanocyte interactions. Exp. Cell Res., 254, 25–32. Lin, C., Chen, N., Scarpa, R., Guan, F., Babiarz-Magee, L., Liebel, F., Li, W., Kizoulis, M., Shapiro, S., and Seiberg, M. (2008) LIGR, a protease-activated receptor-2-derived peptide, enhances skin pigmentation without inducing inﬂammatory processes. Pigment Cell Melanoma Res., 21, 172–183. Derian, C., Eckardt, A., and AndradeGordon, P. (1997) Differential regulation of human keratinocyte growth and differentiation by a novel family of

References

262

263

264

265

266

267

268

269

270

protease-activated receptors. Cell Growth Differ., 8, 743–749. Marthinuss, J., Andrade-Gordon, P., and Seiberg, M. (1995) A secreted serine protease can induce apoptosis in Pam212 keratinocytes. Cell Growth Differ., 6, 807–816. Seiberg, M., Paine, C., Sharlow, E., Andrade-Gordon, P., Constanzo, M., Eisinger, M., and Shapiro, S. (2000) Inhibition of melanosome transfer results in skin lightening. J. Invest. Dermatol., 115, 162–167. Babiarz-Magee, L., Chen, N., Seiberg, M., and Lin, C. (2004) The expression and activation of protease-activated receptor-2 correlate with skin color. Pigment Cell Res., 17, 241–251. Paine, C., Sharlow, E., Liebel, F., Eisinger, M., Shapiro, S., and Seiberg, M. (2001) An alternative approach to depigmentation by soybean extracts via the inhibition of the PAR-2 pathway. J. Invest. Dermatol., 116, 587–595. Zhu, W. and Zhang, R. (2006) Skin lightening agents, in Cosmetic Formulation of Skin Care Products (eds Z.D. Draelos and L.A. Thaman), Cosmetic Science and Technology Series, vol. 30, Taylor & Francis, New York, pp. 205–218. Nair, X., Parah, P., Suhr, L., and Tramposch, K.M. (1993) Combination of 4-hydroxyanisole and all trans retinoic acid produces synergistic skin depigmentation in swine. J. Invest. Dermatol., 101, 145–149. Lei, T., Virador, V., Vieira, W., and Hearing, V. (2002) A melanocyte– keratinocyte coculture model to assess regulators of pigmentation in vitro. Anal. Biochem., 305, 260–268. Berardesca, E., Ardigò, M., Berardesca, M., and Cameli, N. (2008) Melasma: current and future treatments. Expert Rev. Dermatol., 3, 187–193. Kimbrough-Green, C.K., Grifﬁths, C.E., Finkel, L.J., Hamilton, T.A., BulengoRansby, S.M., Ellis, C.N., and Voorhees, J.J. (1994) Topical retinoic acid (tretinoin) for melasma in black patients. A vehicle-controlled clinical trial. Arch. Dermatol., 30, 727–733.

271 Grifﬁths, C.E., Finkel, L.J., Ditre, C.M.,

272

273

274

275

276

277

278

279

280

281

Hamilton, T.A., Ellis, C.N., and Voorhees, J.J. (1993) Topical tretinoin (retinoic acid) improves melasma. A vehicle-controlled, clinical trial. Br. J. Dermatol., 129, 415–421. Leenutaphong, V., Nettakul, A., and Rattanasuwon, P. (1999) Topical isotretinoin for melasma in Thai patients: a vehicle-controlled clinical trial. J. Med. Assoc. Thai., 82, 868–875. Dogra, S., Kanwar, A.J., and Parsad, D. (2002) Adapalene in the treatment of melasma: a preliminary report. J. Dermatol., 29, 539–540. Kasraee, B., Fallahi, M.R., Ardekani, G.S., Ebrahimi, S., Doroudchi, G., Omrani, G.R., Handjani, F., Amini, M., Tanideh, N., Haddadi, M., Nikbakhsh, M., Jahanbani, S., Tran, C., Sorg, O., and Saurat, J.H. (2006) Retinoic acid synergistically enhances the melanocytotoxic and depigmenting effects of monobenzylether of hydroquinone in black guinea pig skin. Exp. Dermatol, 15, 509–514. Kasraee, B., Tran, C., Sorg, O., and Saurat, J.H. (2005) The depigmenting effect of RALGA in C57BL/6 mice. Dermatology, 210 (Suppl. 1), 30–34. Badreshia-Bansal, S. and Draelos, Z. (2007) Insight into skin lightening cosmeceuticals for women of color. J. Drugs Dermatol., 6, 32–39. Cotelessa, C., Peris, K., Onorati, M.T., Fargnoli, M.C., and Chimenti, S. (1999) The use of chemical agents in the treatment of different cutaneous hyperpigmentations. J. Dermatol. Surg., 25, 450–454. Smith, W. (1999) The effect of topical l(+)-lactic acid and ascorbic acid on skin whitening. Int. J. Cosmet. Sci., 21, 33–40. Kubo, I., Kinst-Hori, I., and Yokokawa, Y. (1994) Tyrosinase inhibitors from Anacardium occidentale fruits. J. Nat. Prod., 57, 545–551. Khunger, N., Sarkar, R., and Jain, R.K. (2004) Tretinoin peel versus glycolic peels in the treatment of melasma in dark-skinned patients. Dermatol. Surg., 30, 756–760. Kligman, D.E. (2004) Tretinoin peels versus glycolic peels. Dermatol. Surg., 30, 1609.

163

164

5 Inhibitors and Enhancers of Melanogenesis 282 Halder, R. and Nordlund, J. (2006)

283

284

285

286

287

288

289

Topical treatment of pigmentary disorders, in The Pigmentary System: Physiology and Pathophysiology, 2nd edn (eds J.J. Nordlund, R.E. Boissy, V.J. Hearing, R.A. King, W.S. Oetting, and J.P. Ortonne), Blackwell, Malden, MA, pp. 1165–1174. Yokota, T., Nishio, H., Kubota, Y., and Mizoguchi, M. (1998) The inhibitory effect of glabridin from liquorice extracts on melanogenesis and inﬂammation. Pigment Cell. Res., 11, 355–361. Dorr, R.T., Dvorakova, K., Brooks, C., Lines, R., Levine, N., Schram, K., Miketova, P., Hruby, V., and Alberts, D.S. (2000) Increased eumelanin expression and tanning is induced by a superpotent melanotropin [Nle4-d-Phe7]alpha-MSH in humans. Photochem. Photobiol., 72, 526–532. Barnetson, R.S., Ooi, T.K., Zhuang, L., Halliday, G.M., Reid, C.M., Walker, P.C., Humphrey, S.M., and Kleinig, M.J. (2006) [Nle4-d-Phe7]-alpha-melanocytestimulating hormone signiﬁcantly increased pigmentation and decreased UV damage in fair-skinned Caucasian volunteers. J. Invest. Dermatol., 126, 1869–1878. Fitzgerald, L.M., Fryer, J.L., Dwyer, T., and Humphrey, S.M. (2006) Effect of MELANOTAN, [Nle(4), d-Phe(7)]-alphaMSH, on melanin synthesis in humans with MC1R variant alleles. Peptides, 27, 388–394. Dorr, R.T., Ertl, G., Levine, N., Brooks, C., Bangert, J.L., Powell, M.B., Humphrey, S., and Alberts, D.S. (2004) Effects of a superpotent melanotropic peptide in combination with solar UV radiation on tanning of the skin in human volunteers. Arch. Dermatol., 140, 827–835. Sriwiriyanont, P., Ohuchi, A., Hachiya, A., Visscher, M.O., and Boissy, R.E. (2006) Interaction between stem cell factor and endothelin-1: effects on melanogenesis in human skin xenografts. Lab. Invest., 86, 1115–1125. Wen-Jun, L., Hai-Yan, W., Wei, L., Ke-Yu, W., and Rui-Ming, W. (2008) Evidence that geniposide abrogates norepinephrine-induced

290

291

292

293

294

295

296

297

298

hypopigmentation by the activation of GLP-1R-dependent c-kit receptor signaling in melanocyte. J. Ethnopharmacol., 118, 154–158. Birlea, S.A., Costin, G.E., and Norris, D.A. (2008) Cellular and molecular mechanisms involved in the action of vitamin D analogs targeting vitiligo depigmentation. Curr. Drug Targets, 9, 345–359. Bilodeau, M.L., Greulich, J.D., Hullinger, R.L., Bertolotto, C., Ballotti, R., and Andrisani, O.M. (2001) BMP-2 stimulates tyrosinase gene expression and melanogenesis in differentiated melanocytes. Pigment Cell Res., 14, 328–336. Kawakami, T., Kimura, S., Kawa, Y., Kato, M., Mizoguchi, M., and Soma, Y. (2008) BMP-4 upregulates Kit expression in mouse melanoblasts prior to the Kit-dependent cycle of melanogenesis. J. Invest. Dermatol., 128, 1220–1226. Yaar, M., Wu, C., Park, H.Y., Panova, I., Schutz, G., and Gilchrest, B.A. (2006) Bone morphogenetic protein-4, a novel modulator of melanogenesis. J. Biol. Chem., 281, 25307–25314. Mal’tsev, V.I., Kaliuzhnaia, L.D., and Gubko, L.M. (1995) [Experience in introducing the method of placental therapy in vitiligo in Ukraine]. Lik. Sprava, 40, 123–125. Spry, M.L., Vanover, J.C., Scott, T., Abona-Ama, O., Wakamatsu, K., Ito, S., and D’Orazio, J.A. (2009) Prolonged treatment of fair-skinned mice with topical forskolin causes persistent tanning and UV protection. Pigment Cell Melanoma Res., 22, 219–229. Passeron, T., Namiki, T., Passeron, H.J., Le Pape, E., and Hearing, V.J. (2009) Forskolin protects keratinocytes from UV-B-induced apoptosis and increases DNA repair independent of its effects on melanogenesis. J. Invest. Dermatol., 129, 162–166. Hadshiew, I.M., Eller, M.S., Gasparro, F.P., and Gilchrest, B.A. (2001) Stimulation of melanogenesis by DNA oligonucleotides: effect of size, sequence and 5′ phosphorylation. J. Dermatol. Sci., 25, 127–138. Faas, L., Venkatasamy, R., Hider, R.C., Young, A.R., and Soumyanath, A. (2008)

References In vivo evaluation of piperine and synthetic analogues as potential treatments for vitiligo using a sparsely pigmented mouse model. Br. J. Dermatol., 158, 941–950. 299 Kapoor, R., Phiske, M.M., and Jerajani, H.R. (2009) Evaluation of safety and efﬁcacy of topical prostaglandin E2 in treatment of vitiligo. Br. J. Dermatol., 160, 861–863. 300 Jeon, S., Kim, N.H., Koo, B.S., Lee, H.J., and Lee, A.Y. (2007) Bee venom stimulates human melanocyte proliferation, melanogenesis, dendricity

and migration. Exp. Mol. Med., 39, 603–613. 301 Lee, J.S., Choi, Y.M., and Kang, H.Y. (2007) PPAR-gamma agonist, ciglitazone, increases pigmentation and migration of human melanocytes. Exp. Dermatol., 16, 118–123. 302 Burchill, S.A., Marks, J.M., and Thody, A.J. (1990) Tyrosinase synthesis in different skin types and the effects of alpha-melanocyte-stimulating hormone and cyclic AMP. J. Invest. Dermatol., 95, 558–561.

165

wwwwwww

167

6 Structure of Melanins Shosuke Ito, Kazumasa Wakamatsu, Marco d’Ischia, Alessandra Napolitano, and Alessandro Pezzella

6.1 Introduction

Melanins (Greek μελαηoς = black) is a descriptive term relating to a variety of black, brown, or even yellowish and reddish natural biopolymers of diverse nature and chemical composition that arise biogenetically from the oxidation of phenolic metabolites [1, 2]. The term melanins refers to two main groups of intracellular nitrogenous pigments, the black/dark brown eumelanins (εν = good) and the lighter, yellowish/brown, sulfur-containing pheomelanins (φαεoς = dusky), which arise from a bifurcation of a common biosynthetic pathway involving the tyrosinasecatalyzed oxidation of tyrosine. All too often, however, the term melanins is used synonymously with eumelanins or, more in general, to denote any black insoluble pigment of phenolic origin. In this broader usage, melanins include the pigments found in higher plants, fungi, and bacteria. However, in this chapter we focus exclusively on those pigments derived biogenetically from tyrosine and produced intracellularly within specialized cells. Working on melanins has usually been regarded as an intriguing, although sometimes frustrating, experience. This is due to the extreme heterogeneity of their molecular systems and several adverse properties, including for eumelanins an almost complete insolubility in all solvents, an amorphous particulate character, and the lack of well-deﬁned spectral features. Additional complexities stem from the biological matrix (melanosome) in which the pigments are usually found. Thus, it may not be surprising that melanins’ fundamental structure (if indeed the term “structure” can be applied to such heterogeneous materials) is still uncertain despite extensive studies. Apart from extensive physicochemical investigations, traditional approaches to melanin structure have been based on biosynthetic studies modeling the process of melanogenesis in vitro coupled with chemical degradation leading to fragments of variable diagnostic signiﬁcance. These approaches form the core of this chapter, which is aimed at illustrating current views about melanin structural properties as derived from the biosynthetic and degradative approach. In addition, the main analytical Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

168

6 Structure of Melanins

methods applicable for melanin determination in pigmented tissues are surveyed.

6.2 Classiﬁcation and General Properties of Melanins

Several reviews of melanin structure and chemistry are available to which the interested reader is referred for a more in-depth description of these pigments [2–4]. Although eumelanins and pheomelanins have the primary biogenetic precursor tyrosine in common, they differ signiﬁcantly in their chemical composition and physical properties. Typical eumelanins include the pigments that can be extracted from human and mammalian black/brown hair and irides, as well as those found in the inner ear and melanomas. Eumelanin pigments are also found in various vertebrates, like birds, reptiles, amphibians, and ﬁsh. The most remarkable example of eumelanin from lower animals is represented by the ink of cephalopods, including Sepia ofﬁcinalis, Octopus vulgaris, and Loligo vulgaris. Studies of natural eumelanins have been carried out mainly on pigments isolated from Sepia ink, black human hair, and melanoma specimens. Isolation procedures are basically aimed at removing the protein matrix of the tissue while limiting oxidative damage to the pigment. A typical method for isolating eumelanin from black human hair involves sequential digestion with protease, proteinase K, and papaine in the presence of dithiothreitol, leading to a pigment with a 6–14% (w/w) protein content [5, 6]. Eumelanins display a peculiar set of physicochemical properties that have been considered an operational deﬁnition of eumelanins [7]. In the condensed phase or ﬁnely dispersed in an aqueous medium most eumelanins exhibit a featureless broadband monotonic absorption accounting for their black appearance. The origin of the broad spectrum of eumelanin is attributed to different contributions, including absorption and scattering. Electrical conductivity and photoconductivity are other peculiar properties that distinguish eumelanins from other bio-organic materials. Despite intensive investigations, the origin of these properties and the mechanism by which a charge is generated and transported are still poorly understood. Eumelanins are capable of reducing and oxidizing other molecules – a property that stems from the redox behavior of its monomer units – and display a persistent electron paramagnetic resonance (EPR) signal indicating free radical centers. The ability to bind various metal ions is still another characteristic feature of eumelanins (e.g., [8]). Whereas eumelanins are relatively widespread in nature, pheomelanins are found only in mammalian skin, hair, and eyes, and in bird feathers [9, 10]. In mice and other mammals, pheomelanin in the typical agouti-banding pattern provides camouﬂage. Natural pheomelanins are difﬁcult to isolate from their biological matrix and are usually obtained with a high protein content [6]. The most charac-

6.3 Biosynthetic Studies

teristic feature of pheomelanins is their sulfur content, which accounts for up to around 10% in pigment samples from red feathers and is due to the incorporation of cysteine in the biosynthetic pathway [4, 11]. Most of the studies of pheomelanins have been done on partially puriﬁed samples obtained from hen feathers (gallopheomelanins), red human hair, and mammalian fur. Puriﬁcation protocols should avoid harsh treatments causing degradative changes to the pheomelanin and should preferably involve repeated enzymatic digestions [6, 12]. Unlike eumelanins, which are completely insoluble in most solvents, pheomelanins exhibit a limited solubility in aqueous alkali. Natural and synthetic pheomelanins display remarkably similar absorption spectra featuring a ﬂat maximum around 305 nm, and inﬂections at around 260 and 360 nm, with limited absorption above 450– 500 nm [13]. Apparently different molecular constituents in pheomelanin are responsible for the chromophoric features. The photoreactive chromophores of pheomelanins are of low molecular weight and exhibit similar photophysics in the aggregated state. They may include benzothiazine structural motifs [14], but there appears to be also a signiﬁcant participation of benzothiazoles and other structural units of the trichochrome type [13, 15]. Pheomelanins exhibit an EPR signal with distinct immobilized nitroxide-like features attributed to a partial localization of the unpaired electron on the nitrogen atom of the o-semiquinone imine form of 1,4-benzothiazine subunits [16]. Like eumelanins, pheomelanins also bind metal cations, with a Fe(III) content that is 4 times higher in the red pheomelanosomes as compared with the black eumelanosomes [6].

6.3 Biosynthetic Studies 6.3.1 Early Stages of Melanogenesis

Eumelanin and pheomelanin are both derived from the common precursor dopaquinone formed by oxidation of the common amino acid l-tyrosine by tyrosinase (Scheme 6.1; see also Chapters 3 and 4). Dopaquinone is the ﬁrst intermediate during the initial stage of melanogenesis [17]. As an o-quinone, dopaquinone is highly reactive and in the absence of sulfydryl compounds it undergoes the intramolecular addition of the amino group to produce cyclodopa (also called leucodopachrome). The chemistry of o-quinones is described in detail in Chapter 3, and Ito and Wakamatsu [18] also summarize pivotal roles of dopaquinone in controlling melanogenesis. A redox exchange between cyclodopa and dopaquinone then gives rise to dopachrome, an orange/red intermediate, and 3,4-dihydroxyphenylalanine (dopa). This latter reaction is considered the source of dopa formed during melanogenesis (Scheme 6.1). Dopachrome then gradually rearranges to generate mostly 5,6-dihydroxyindole (DHI) and to a minor extent DHI-2-carboxylic acid (DHICA) [19]. Under spontaneous reaction at neutral pH,

169

170

6 Structure of Melanins

COOH NH2

HO

tyrosine Tyrosinase

HO

O2 Tyrosinase

DOPA

COOH Cys NH2

O

COOH NH2

HO

O2

O

COOH

HO

NH2

HO S

dopaquinone

H2N COOH cysteinyldopa O O

HO

N H cyclodopa

N H DHI [O]

HO

EUMELANIN

N

OH NH2

S

HO

HO

COOH

(HOOC)

tyrp-2

–CO2

HO

HO

COOH N H dopachrome

N H DHICA [O]

COOH

COOH

[O] PHEOMELANIN

OH N

(COOH)H N 2 COOH S

S HOOC

NH2

O

N H

OH

TRICHOCHROMES Schematic outline of the melanogenetic pathway showing the switch from eumelanin to pheomelanin by intervention of cysteine.

Scheme 6.1

DHI and DHICA are formed in a ratio of 70 : 1 [20]. These two DHIs are then further oxidized and polymerized to produce eumelanin. In addition to tyrosinase, two tyrosinase-related proteins have been shown to modulate/accelerate eumelanogenesis (see Chapter 4 for more details). Dopachrome tautomerase (Dct; also known as tyrosinase-related protein 2 (TRP-2/ Tyrp2)) catalyzes the tautomerization of dopachrome to DHICA [21, 22]. Certain divalent metal ions, especially Cu2+, can also promote tautomerization of dopachrome to DHICA, but Dct seems to be more effective in catalyzing the tautomerization [20, 23]. The oxidative polymerization of DHI is catalyzed by mammalian tyrosinase [24]. A recent pulse radiolysis study, however, indicates that DHI can be effectively oxidized by dopaquinone through redox exchange [25]. On the other hand, oxidative polymerization of DHICA appears to be catalyzed by tyrosinase in humans [26] or by Tyrp1 in mice [27, 28]. Thus, the activities of these tyrosinase-related

6.3 Biosynthetic Studies

proteins greatly affect the quantity and quality (the ratio of DHI to DHICA and the degree of polymerization) of eumelanins produced. Regarding the production of pheomelanin, the intervention of sulfydryl compounds such as cysteine gives rise exclusively to thiol adducts of dopa, cysteinyldopas. Tyrosinase oxidation of dopa in the presence of excess cysteine produces a high yield of 5-S-cysteinyldopa (74%) and 2-S-cysteinyldopa (14%) together with minor amounts of 6-S-cysteinyldopa (1%) and a di-adduct, 2,5-S,S′-dicysteinyldopa (5%) [29]. Further oxidation of the cysteine adducts leads to the formation of pheomelanin via benzothiazine intermediates [14]. 6.3.2 Late Stages of Eumelanogenesis

Based on elemental analysis, chemical degradation, and isotope labeling experiments [19, 30, 31], it has been concluded that the proportion of DHI- and DHICAderived units in eumelanins varies signiﬁcantly depending on the origin of the pigment (natural or synthetic); in particular, intact natural eumelanins contain a higher proportion of DHICA (up to 75%) than enzymatically prepared synthetic eumelanins (less than 10%). Investigation of the oxidative polymerization of DHIs has been used as a strategy for inquiring into the mode of formation of eumelanins as well as for developing a consistent structural model [32]. The role of DHI and DHICA as the basic eumelanin building blocks is indicated by their rapid conversion to black insoluble pigments following exposure to oxidizing enzymes, UV radiation, chemical oxidants, or even on standing at neutral physiologic pH. Oxidation of DHI leads to a collection of dimers and trimers that reveals the prevalent mode of coupling of the indole monomer through 2,4′- and 2,7′bondings (Scheme 6.2). A uniﬁed synthetic strategy for the preparation of 2,7′-, 2,2′-, and 2,3′-dimers has recently been developed [33]. The mechanism of indole dimerization is uncertain, but a catechol–quinone interaction seems the most likely route based on product structures, theoretical calculations [34], and quinone trapping experiments with sulfur nucleophiles [35]. Pulse radiolysis experiments suggest that one-electron oxidation of DHI in aqueous solution at pH 7.4 leads to a semiquinone, which decays by a second-order process [25, 36]. Oxidative coupling of dimers proceeds likewise via semiquinone and quinone intermediates [37], and leads to tetramers in which other types of interring bonds are formed (e.g., 2,3′-, 4,4′-, and 7,7′-bonds), depending on the dimer substrate [38, 39]. These model studies point to a mechanism of polymerization of DHI in which monomer units couple through the 2- and 4- or 7-positions. However, as oligomer coupling steps become signiﬁcant, other modes of bonding may become important, with consequent bending of the growing oligomer chain. As a result, a high degree of structural diversity is expectedly generated during eumelanin biosynthesis, accounting for the marked heterogeneity of the pigment. It may be relevant in this regard to mention the formation of a cyclic pentamer by co-oxidation of a dimer and a trimer from 5,6-dihydroxy-1-methylindole [40],

171

172

6 Structure of Melanins

HO HO [O] HO

HO

OH

HO

NH +

N H HN

HO

[O] HO metal cations N HO H

HO

DHI N HHO

HO

OH

HO OH N H HN

HO HO HO

HO HO

H HN N

HO

+ HO HO NH

N H HO

OH HO N NH H

OH

OH OH

+

OH

NH H N

N H HN

HO

OH

OH

OH H N

HN

N H

OH

OH

HO

HO

HO

+ HO

N H

HO OH

N H NH

HO OH HO

NH

OHHO HO

HO

Scheme 6.2

HO

N H

OH

[O]

[O] HO

OH

N H HO

OH

NH

N H

OH N H

OH

OH

Main oligomers formed by oxidation of DHI.

raising the possibility that cyclic structural motifs are involved in the process of eumelanin formation [41]. Oxidation of DHICA is a process of considerable biological importance considering not only the prevalent involvement of this indole in the biosynthesis of natural eumelanins [19] and ocular pigments (e.g., the brown tapetum lucidum of the sea catﬁsh (Arius felis L.) [42] ), but also its putative role as an endogenous antioxidant [43, 44]. Since the carboxylic group at C2 limits the range of options available for indole coupling, a lower number of positional isomers are expected to populate the various oligomer stages. Main oligomers formed by oxidative coupling of DHICA include the 4,4′-biindolyl, the 4,7′-biindolyl, and other minor dimers, a series of trimers, and tetramers [32, 45, 46] (Scheme 6.3). Notably, oligomers of DHICA exhibit hindered rotation around inter-ring bonds and the negatively charged carboxylate functions contribute to maintain signiﬁcant twist angles between units. It is therefore suggested that the effective conjugation length in oligomeric/polymeric eumelanin components from DHICA may be controlled by such hindered rotation around interring bonds preventing planarization of the continuous array of indole units [47]. This may provide an explanation for the difference in the absorption properties of polymers from the two key eumelanin monomers.

6.3 Biosynthetic Studies

HO CO2H HO

N H [O]

HO

HO

CO2H

N H CO2H

HO HO

HO

N H

HO

N H

N H

HO

HO OH HN

OH

HO

HO2C HO HO

N H

CO2H

H N

CO2H

HO N H

HO HO

N H

CO2H

CO2H

CO2H

HO CO2H

CO2H N H [O]

CO2H

H N

HO N H N H

HO N H

CO2H

N H H N

CO2H

CO2H

HO HO

CO2H

CO2H

HO

HO

HO

HO

CO2H

HO

HO

HO

H N

HO HO

[O]

HO

HO HO

HO

N H H N

HO HO

HO DHICA

HO

HO

CO2H

HO

CO2H N H

CO2H

Scheme 6.3 Main oligomers formed by oxidation of DHICA.

Despite the extensive investigations described above, the detailed structural features of DHI polymers are largely unknown. To what extent DHI and DHICA oligomers model eumelanin fundamental building blocks is a most critical issue, and attention is also called to the possibility that DHI and DHICA can form copolymeric structures on oxidation [48, 49]. Early structural models envisaged a high-molecular-weight heteropolymeric system formed via random bonding between different DHI units. Later, an alternative view of eumelanin structure was proposed, based on scanning tunneling microscopy and X-ray studies [50–52], which suggested that eumelanins were not large extended heteropolymers, but rather mixtures of π-stacked oligomeric “protomolecules” consisting of three to four planar layers of no more than ﬁve or six indolic units and 15–20 Å in size. It should be emphasized, however, that this

173

174

6 Structure of Melanins

model would apply only to planar oligomer sheets made up of DHI units and not to the twisted DHICA oligomers, and is based on hypothetical structures devoid of experimental support. In this regard, small-angle X-ray scattering (SAXS) data [53] and atomic force microscopy (AFM) investigations of sepia eumelanin [54] would be compatible with eumelanins being mixtures of oligomers. Matrixassisted laser desorption/ionization (MALDI) mass spectroscopic studies have also identiﬁed several oligomeric and partially degraded species both in sepia melanin and synthetic eumelanins [55]. The structural and electronic properties of putative monomer and oligomer components of eumelanin architecture have also been investigated by theoretical methods, showing that different oxidation states and tautomeric forms can have dramatically different highest occupied molecular orbital to lowest unoccupied molecular orbital (HOMO–LUMO) gaps, and that an increase in oligomer size and π-stacking induce a signiﬁcant compression of the gap, with bathochromic shift of the absorption maximum [56, 57]. The concepts emerging from theoretical studies seem to lend support to oligomer-based structures in which a broad range of variations is possible in terms of molecular size and redox states. However, further work is needed before the structural problem is deﬁnitively settled. As mentioned earlier, a fundamental issue concerning eumelanins is why they are black. The experimental difﬁculty lies in the virtual insolubility of these pigments, and the associated scattering effects and hindering characterization of the intrinsic absorption properties of the heterogeneous species produced by oxidative polymerization of monomer precursors. Important insights into this issue have derived from studies of soluble eumelanin-like materials aimed at disentangling intrinsic absorption properties of basic components by circumventing scattering effects. This goal was recently achieved, for example, by polymerizing DHI in the presence of polyvinylalcohol [58] and by oxidation of a glycated derivative of DHI (5,6-dihydroxy-3-indolyl-1-thio-β-d-galactopyranoside) [59]. The resulting dark brown soluble polymer exhibited a distinct UV band around 315 nm and a broad visible absorption, resembling that of natural eumelanins. By a systematic series of reduction and dilution experiments it was established that eumelanin black color is not only intrinsically deﬁned by the overlap of π-electron conjugated chromophores within the individual polymer components, as commonly believed, but also by oxidation state- and aggregation-dependent interchromophoric interactions causing perturbations of the heterogeneous ensemble of π-electron systems and overall spectral broadening. These ﬁndings may overall set the basis for an improved interpretative model of eumelanin structure and properties that is based on a dynamic molecular disorder with extensive intermolecular and intramolecular redox and charge transfer interactions. 6.3.3 Late Stages of Pheomelanogenesis

Oxidation of dopa in the presence of cysteine is undoubtedly the process that mimics at best pheomelanin synthesis within pigment cells. However, oxidation

6.3 Biosynthetic Studies

of 5-S-cysteinyldopa apparently leads to less heterogeneous pigments that may be more useful for structural investigations. The reaction involves the formation of an o-quinone, which rapidly cyclizes to give an unstable o-quinonimine intermediate. This may undergo redox exchange with cysteinyldopa leading to a dihydrobenzothiazine intermediate, or may be converted to 2H-1,4-benzothiazine species by rearrangement with (85%) or without (15%) decarboxylation [11, 60]. These may be oxidized to pheomelanins by way of different processes, including phenolic dimerizations [61] and imine–enamine reactions with coupling at the 2-position to give trichochrome-like dimers [62, 63]. Notably, small amounts of trichochromelike pigments are slowly formed by oxidation of cysteinyldopas, a process speeded up by acidiﬁcation of the reaction mixtures [64], but whether this behavior has some bearing on the identiﬁcation of trichochromes in pigmented tissues remains to be assessed. Oxygenated systems related to 3-oxo-2H-1,4-benzothiazine units and benzothiazole moieties are also likely to be involved in pigment formation (Scheme 6.4). A recent biosynthetic study starting from dopa and cysteine [14] has shown that dihydro-1,4-benzothiazine species are the last major, isolable intermediates in pheomelanogenesis, and are then oxidized and polymerized to give pheomelanin with a gradual conversion of the benzothiazine units to benzothiazole units in polymer backbone. Beyond this level, knowledge of the molecular mechanism of pheomelanogenesis is virtually lacking. Nevertheless, it should be stressed that dopaquinone undergoes redox exchange reactions with cysteinyldopas and dihydro-1,4-benzothiazines to generate the corresponding o-quinone or o-quinonimine, thus spontaneously promoting pheomelanogenesis [14]. It stresses again the pivotal roles of dopaquinone in promoting the whole process of (mixed) melanogenesis. 6.3.4 Concept of Mixed Melanogenesis

Kinetic studies using the pulse radiolysis technique have provided a number of useful insights into the dynamic process of mixed melanogenesis. Thus, melanogenesis proceeds in three distinct stages (see [18] for more detail). The initial stage is the production of cysteinyldopas, as long as cysteine concentration is above 0.13 μM. The second stage is the oxidation of cysteinyldopas by dopaquinone to produce pheomelanins, which continues to proceed as long as cysteinyldopas are present at concentrations above 9 μM. The last stage is the production of eumelanins, which begins only after most cysteinyldopas (and cysteine) are depleted. Therefore, the ratio of eumelanin to pheomelanin is determined by tyrosinase activity, and the availability of tyrosine and cysteine in melanosomes [65]. This time course of melanogenesis further suggests the “casing model” for mixed melanogenesis. Thus, in the process of mixed melanogenesis, pheomelanic pigment is produced ﬁrst making a core, followed by the deposit of eumelanic pigment to make an outer layer [18]. This model was originally proposed by Agrup et al. [66], based on biochemical ﬁndings, and then Bush et al. [67] provided direct, biophysical evidence that supports this model, by showing that neuromelanin

175

176

6 Structure of Melanins

HOOC

OH

HOOC NH2

NH2

OH

5-S-cysteinyldopa HOOC

O O S

S HOOC

NH2

NH2

O

HOOC OH NH2 HOOC

NH2

H N

N

(COOH)

S

COOH

S OH NH2 HOOC OH

NH2 HOOC

H N S

N

(COOH) OH

S

N

(COOH) H N 2

S

O HOOC OH

3-oxo-3,4-dihydrobenzothiazine N NH2 (λ max 299 nm) S HOOC benzothiazoles

NH2 (HOOC)

COOH

S N

OH 2,2'-bi(2H-1,4-benzothiazine) (COOH) (λ max 346 nm )

PHEOMELANIN Oxidation routes of 5-S-cysteinyldopa to pheomelanin through benzothiazine intermediates. Scheme 6.4

granules have a eumelanic surface with a pheomelanic core. The casing model was further supported on mixed melanins produced in iridal melanocytes [4, 68]. Although those melanins have varying ratios of eumelanin to pheomelanin, they have the surface oxidation potentials characteristic for eumelanin.

6.4 Degradative Studies 6.4.1 Eumelanins

Owing to their high chemical heterogeneity, elemental analyses of eumelanins lack reproducibility. The data vary signiﬁcantly with the nature of the sample, isola-

6.4 Degradative Studies

HO HO

HO

HO COOH

N H

N H

HO

2-substituted DHI units

DHICA units

[O]

2-unsubstituted DHI units [O]

[O] HOOC

HOOC HOOC

N H

HO

N H

COOH

pyrrole-2,3,5-tricarboxylic acid (PTCA)

HOOC

N H

pyrrole-2,3-dicarboxylic acid (PDCA)

Scheme 6.5 Origin of pyrrole carboxylic acids by oxidative degradation of eumelanins.

tion procedure, storage method, and other experimental parameters, including hydration [1]. However, the C/N ratio provides an indication of the proportion of DHICA versus DHI units [19], with the caveat that it may provide limited information in the case of extensive degradation of the benzene moieties of the indole units and is applicable mainly for synthetic eumelanins. The carboxyl group attached to the indole or pyrrole ring is labile and may easily be split off as CO2 by heating a melanin suspension in 6 M HCl [2, 19]. This methodology has been applied successfully to estimate the carboxyl content in natural and synthetic eumelanins [5, 19, 31, 69]. Chemical analysis of eumelanins is based on oxidative degradation with potassium permanganate [70, 71] or alkaline hydrogen peroxide [72, 73] followed by high-performance liquid chromatography (HPLC) detection of pyrrole-2,3,5tricarboxylic acid (PTCA) and pyrrole-2,3-dicarboxylic acid (PDCA) (Scheme 6.5). In addition to these major degradation products, related pyrrole carboxylic acids such as 2-carboxymethylpyrrole-3,5-dicarboxylic acid are produced [74]. Although usually formed in rather small amounts (below 6–7%), pyrrole carboxylic acids are diagnostic markers for eumelanins and provide perhaps the most cogent evidence for the presence of indolic building blocks in the eumelanin backbone. DHICA-rich polymers give higher yields of PTCA relative to DHI polymers, which in turn furnish more PDCA [71]. DHI units with 2- and 3-positions unsubstituted in eumelanins afford PDCA [75]. Thus, the ratio of PTCA to PDCA may reﬂect the ratio of DHICA-derived units to DHI-derived units in eumelanins. Upon oxidation, pyrrole carboxylic acids arise not only from indolic building blocks but also from degraded pyrrolic building blocks. The (dihydroxy)indolic

177

178

6 Structure of Melanins

OH

OH N

NH2 HOOC

(COOH) HI

S

NH2

NH2 HOOC

pheomelanin structural unit

aminohydroxyphenylalanine (AHP)

oxidative degradation OH NH2 HOOC

N COOH S

benzothiazolecarboxylic acid (BTCA) Scheme 6.6

HOOC

N

HOOC

S

COOH

+

thiazole-2,4,5-tricarboxylic acid (TTCA)

Main degradation products of pheomelanins.

units can provide the redox property of eumelanins while the pyrrolic units cannot. Thus, the ratio of indolic units to pyrrolic units is important in determining the degree of the redox property of eumelanins. In this connection, sepia melanin is highly degraded to a pyrrolic structure, thus implying little redox property compared with the high redox property of synthetic DHI or DHICA melanin [31]. Pyrrole carboxylic acids PTCA and PDCA are present also in the free, extractable form in melanosomes obtained from Sepia, human black hair, and bovine eyes [76]. The free form consists of as much as 20% of the total amounts obtained after hydrogen peroxide oxidation and demonstrates extensive oxidative breakdown of indolic units in natural eumelanins. This degradation is likely to proceed through peroxidative cleavage as occurs in the chemical degradation. In this connection, it should be noted that UV-A irradiation induces ﬂuorescence of melanin accompanied by pigment bleaching [77]. As PTCA and PDCA are ﬂuorescent compounds this provides evidence of degradation by photo-oxidation. 6.4.2 Pheomelanins

Pheomelanin treated with HI leads to the formation of aminohydroxyphenylalanine (AHP) isomers by reductive hydrolysis of benzothiazine moieties (Scheme 6.6). An advantage of hydriodic acid (HI) hydrolysis is the rather high yield (around 20%) of AHP isomers [70, 71, 78]. AHP isomers, 4-amino-3-hydroxyphenylalanine (4-AHP) and 3-amino-4-hydroxyphenylalanine (3-AHP), derive from 5-S-cysteinyldopa- and 2-S-cysteinyldopa-derived benzothiazine units in pheomelanins, respectively [78]. It should be noted that AHP isomers are not derived from benzothiazole

6.4 Degradative Studies

units in the pheomelanin structure [14]. 3-AHP may also arise from 3-nitrotyrosine produced in vivo by nitric oxide-dependent nitration of tyrosine. An alternative approach involves oxidative degradation with alkaline hydrogen peroxide leading to a number of diagnostic structural fragments including thiazole-2,4,5-tricarboxylic acid (TTCA) and thiazole-4,5-dicarboxylic acid (TDCA [79]), while alkaline permanganate produces a series of pyridine-containing polycarboxylic acids, including pyridine-2,3,4,6-tetracarboxylic acid and 2-(2′-(4′,5′-dicarboxythiazolyl))-3,4,6-pyridinetricarboxylic acid [80]. More recent work on human red hair pheomelanin has led to the identiﬁcation of two additional degradation products: the isomeric benzothiazolecarboxylic acids BTCA (from 5-S-cysteinyldopa) and BTCA-2 (from 2-S-cysteinyldopa) [81]. Analysis of different segments of red hair locks (4–20 cm) with the improved dual-marker methodology revealed a remarkable degradation of the 5-Scysteinyldopa-derived constituents on passing from the root region near the scalp (or very short hair) to the tip, whereas 2-S-cysteinyldopa-related units were little modiﬁed throughout the entire hair length. Lethal yellow and recessive yellow mouse hair and red chicken feathers gave values of BTCA and BTCA-2 similar to those of red hair regions near the root, and better reﬂective of the typical 5-Scysteinyldopa/2-S-cysteinyldopa formation ratio. Prolonged exposure of red hair locks to intense sunlight caused a modest decrease in the yields of BTCA, but not BTCA-2. It appears that red hair pheomelanin consists of stable structural components made up of 2-S-cysteinyldopa-derived units, associated to a degradable 5-S-cysteinyldopa component. Degradation occurs during hair growth probably as a result of oxidative processes related in part, but not exclusively, to sun exposure. The structural origin of the isomeric benzothiazole carboxylic acids BTCA and BTCA-2 appears to be benzothiazine units despite of the benzothiazole structure of BTCA and BTCA-2 [80]. Overall, these data have led to the hypothesis that pheomelanins consist of benzothiazine, benzothiazole, and isoquinoline units linked at various levels and through different bondings. However, while benzothiazine and benzothiazole units have been identiﬁed in all natural and synthetic pheomelanins, and have been substantiated by chemical isolation and degradation studies, and a straightforward synthetic access is available [82], the actual presence of isoquinoline units still needs to be demonstrated. The origin of benzothiazole fragments is clearly via contraction of benzothiazine ring precursors, but whether this occurs exclusively during biosynthesis, as has been shown recently [14], or is also in part associated with postsynthetic oxidative degradation processes is still uncertain and requires further elaboration. Speciﬁcally, it should be clariﬁed what type of chemical modiﬁcation takes place during the selective (photo)degradation of 5-S-cysteinyldopa-derived benzothiazine units in pheomelanin, whether it is the ring contraction of benzothiazine to benzothiazole structure or a more complex modiﬁcation of the whole pheomelanin skeleton. A hint to the possible occurrence of the ring contraction to benzothiazole units is the facile conversion of 1,4-dihydrobenzothiazine-3carboxylic acid to 2-methylbenzothiazole upon UV-A radiation [83]. The ring contraction is also markedly promoted by the presence of Fe3+ ions [84].

179

180

6 Structure of Melanins

6.5 Analysis of Eumelanins and Pheomelanins

Regulation of melanogenesis has been a subject of extensive studies. In most such studies, quantiﬁcation of melanins in pigmented tissues such as hair, skin, melanocytes, and melanomas is essential. However, previous methods for characterizing melanins in pigmented tissues required the isolation of melanin pigments. Moreover, none of those methods was suitable for distinguishing eumelanins and pheomelanins [2]. Only one exception is the EPR spectrum in which eumelanins give a single peak while pheomelanins afford two characteristic peaks [16]. Under these circumstances, Ito and Fujita [70] introduced a rapid and sensitive method for quantitatively analyzing eumelanin and pheomelanin in tissue samples by taking advantage of degradative studies. The method is based on the formation of PTCA (Scheme 6.5) by acidic permanganate oxidation of eumelanins and of AHP isomers by HI hydrolysis of pheomelanins. These speciﬁc degradation products are determined by HPLC with UV or electrochemical detection, respectively. An advantage of PTCA by permanganate oxidation as a marker for eumelanins is that it is highly speciﬁc for eumelanins [70, 85]. However, the permanganate oxidation method is rather tedious and not suitable as routine assays, and is now being replaced by alkaline hydrogen peroxide oxidation that is much easier to perform (because of the exclusion of ether extraction and the direct injection into HPLC), and gives higher yields of PTCA and PDCA [72, 73, 86]. The ratio of PTCA to PDCA can be used to assess the ratio of DHICA-derived units to DHI-derived units in eumelanins (unpublished results by Ito and Wakamatsu). The ratios are much higher in mouse hairs than in human hairs, indicating a large difference in the ratio of DHICA to DHI between the two species. Previously, the relative content of DHICA in eumelanins was assessed indirectly from the ratio of PTCA to total melanin [85, 87]. Regarding differential roles of DHICA and DHI as eumelanin building blocks, it should be stressed that DHICA melanin has a high ability to scavenge hydroxyl radicals while DHI melanin does not [44]. However, whether this advantageous property of DHICA unit in eumelanins plays a physiological role in vivo remains to be clariﬁed.

6.6 Conclusions

Melanin pigments, eumelanins and pheomelanins, can be described today as collections of molecular species with different chemical structures and properties, thus encompassing diverse degrees of chemical heterogeneity. This molecular heterogeneity can be ascribed to a number of factors, including: the range of different monomer units (several precursor compounds can take part in the polymerization process); the different modes of inter-unit coupling; differences in molecular weight (arising from the variable degree of polymerization); the redox

References

state; and the extent and mode of aggregation. On this basis, it should be clear that the structure of melanins – even if the term “structure” can rightly be applied to such complex and heterogeneous biomaterials – is a challenging issue that can be addressed only through an integrated spectral, chemical, and degradative approach.

References 1 Prota, G. (1992) Melanins and

2

3

4

5

6

7

8

9

Melanogenesis, Academic Press, San Diego, CA. Ito, S. and Wakamatsu, K. (2006) Chemistry of melanins, in The Pigmentary Systems: Physiology and Pathophysiology (eds J.J. Nordlund, R.E. Boissy, V.J. Hearing, R.A. King, W.S. Oetting, and J.P. Ortonne), 2nd edn, Blackwell, Malden, MA, pp. 282–310. Simon, J.D., Hong, L., and Peles, D.N. (2008) Insights into melanosomes and melanin from some interesting spatial and temporal properties. J. Phys. Chem. B, 112, 13201–13217. Simon, J.D., Peles, D.N., Wakamatsu, K., and Ito, S. (2009) Current challenges in understanding melanogenesis: bridging chemistry, biological control, morphology, and function. Pigment Cell Melanoma Res., 22, 563–579. Novellino, L., Napolitano, A., and Prota, G. (2000) Isolation and characterization of mammalian eumelanins from hair and irides. Biochim. Biophys. Acta, 1475, 295–306. Liu, Y., Hong, L., Wakamatsu, K., Ito, S., Adhyaru, B., Cheng, C.Y., Bowers, C.R., and Simon, J.D. (2005) Comparison of structural and chemical properties of black and red human hair melanosomes. Photochem. Photobiol., 81, 135–144. Meredith, P. and Sarna, T. (2006) The physical and chemical properties of eumelanin. Pigment Cell Res., 19, 572–594. Liu, Y. and Simon, J.D. (2005) Metal-ion interactions and the structural organization of Sepia eumelanin. Pigment Cell Res., 18, 42–48. Ito, S. and Wakamatsu, K. (2003) Quantitative analysis of eumelanin and pheomelanin in humans, mice, and other

10

11

12

13

14

15

16

animals: a comparative review. Pigment Cell Res., 16, 523–531. McGraw, K. (2008) An update on the honesty of melanin-based color signals in birds. Pigment Cell Melanoma Res, 21, 133–138. Di Donato, P. and Napolitano, A. (2003) 1,4-Benzothiazines as key intermediates in the biosynthesis of red hair pigment pheomelanins. Pigment Cell Res., 16, 532–539. Panzella, L., Manini, P., Monfrecola, G., d’Ischia, M., and Napolitano, A. (2007) An easy-to-run method for routine analysis of eumelanin and pheomelanin in pigmented tissues. Pigment Cell Res., 20, 128–133. Napolitano, A., De Lucia, M., Panzella, L., and d’Ischia, M. (2008) The “benzothiazine” chromophore of pheomelanins: a reassessment. Photochem. Photobiol., 84, 593–599. Wakamatsu, K., Ohtara, K., and Ito, S. (2009) Chemical analysis of late stages of phaeomelanogenesis: conversion of dihydrobenzothiazine to a benzothiazine structure. Pigment Cell Melanoma Res., 22, 474–486. Ye, T., Pawlak, A., Sarna, T., and Simon, J.D. (2008) Different molecular constituents in pheomelanin are responsible for emission, transient absorption and oxygen photoconsumption. Photochem. Photobiol., 84, 437–443. Sealy, R.C., Hyde, J.S., Felix, C.C., Menon, I.A., Prota, G., Swartz, H.M., Persad, S., and Haberman, H.F. (1982) Novel free radicals in synthetic and natural pheomelanins: distinction between dopa melanins and cysteinyldopa melanins by ESR spectroscopy. Proc. Natl. Acad. Sci. USA, 79, 2885–2889.

181

182

6 Structure of Melanins 17 Cooksey, C.J., Garratt, P.J., Land, E.J.,

18

19

20

21

22

23

24

25

26

Pavel, S., C.A. Ramsden, Riley, P.A., and Smit, N.P. (1997) Evidence of the indirect formation of the catecholic intermediate substrate responsible for the autoactivation kinetics of tyrosinase. J. Biol. Chem., 272, 26226–26235. Ito, S. and Wakamatsu, K. (2008) Chemistry of mixed melanogenesis – pivotal roles of dopaquinone. Photochem. Photobiol., 84, 582–592. Ito, S. (1986) Reexamination of the structure of eumelanin. Biochim. Biophys. Acta, 883, 155–161. Palumbo, A., d’Ischia, M., Misuraca, G., and Prota, G. (1987) Effect of metal ions on the rearrangement of dopachrome. Biochim. Biophys. Acta, 925, 203–209. Pawelek, J., Körner, A., Bergstrom, A., and Bologna, J. (1980) New regulators of melanin biosynthesis and the autodestruction of melanoma cells. Nature, 286, 617–619. Tsukamoto, K., Jackson, I.J., Urabe, K., Montague, P.M., and Hearing, V.J. (1992) A second tyrosinase-related protein, TRP-2, is a melanogenic enzyme termed DOPAchrome tautomerase. EMBO J., 11, 519–526. Palumbo, A., Solano, F., Misuraca, G., Aroca, P., García-Borròn, J.C., Lozano, J.A., and Prota, G. (1991) Comparative action of dopachrome tautomerase and metal ions on the rearrangement of dopachrome. Biochim. Biophys. Acta, 1115, 1–5. Tripathi, R.K., Hearing, V.J., Urabe, K., Aroca, P., and Spritz, R.A. (1992) Mutational mapping of the catalytic activities of human tyrosinase. J. Biol. Chem., 267, 23707–23712. Edge, R., d’Ischia, M., Land, E.J., Napolitano, A., Navaratnam, S., Panzella, L., Pezzella, A., Ramsden, C.A., and Riley, P.A. (2006) Dopaquinone redox exchange with dihydroxyindole and dihydroxyindole carboxylic acid. Pigment Cell Res., 19, 443–450. Olivares, C., Jiménez-Cervantes, C., Lozano, J.A., Solano, F., and GarcíaBorrón, J.C. (2001) The 5,6-dihydroxyindole-2-carboxylic acid

27

28

29

30

31

32

33

34

(DHICA) oxidase activity of human tyrosinase. Biochem. J., 354, 131–139. Jiménez-Cervantes, C., Solano, F., Kobayashi, T., Urabe, K., Hearing, V.J., Lozano, J.A., and García-Borrón, J.C. (1994) A new enzymatic function in the melanogenic pathway. The 5,6-dihydroxyindole-2-carboxylic acid oxidase activity of tyrosinase-related protein-1. J. Biol. Chem., 269, 17993–18000. Kobayashi, T., Urabe, K., Winder, A.J., Jiménez-Cervantes, C., Imokawa, G., Brewington, T., Solano, F., GarcíaBorrón, J.C., and Hearing, V.J. (1994) Tyrosinase related protein 1 (TRP1) functions as a DHICA oxidase in melanin biosynthesis. EMBO J., 13, 5818–5825. Ito, S. and Prota, G. (1977) A facile one-step synthesis of cysteinyldopas using mushroom tyrosinase. Experientia, 33, 1118–1119. Tsukamoto, K., Palumbo, A., d’Ischia, M., Hearing, V.J., and Prota, G. (1992) 5,6-Dihydroxyindole-2-carboxylic acid is incorporated in mammalian melanin. Biochem. J., 286, 491–495. Pezzella, A., d’Ischia, M., Napolitano, A., Palumbo, A., and Prota, G. (1997) An integrated approach to the structure of Sepia melanin. Evidence for a high proportion of degraded 5,6-dihydroxyindole-2-carboxylic acid units in the pigment backbone. Tetrahedron, 53, 8281–8286. d’Ischia, M., Napolitano, A., Pezzella, A., Land, E.J., Ramsden, C.A., and Riley, P.A. (2005) 5,6-Dihydroxyindoles and indole-5,6-diones. Adv. Heterocycl. Chem., 89, 1–63. Capelli, L., Manini, P., Pezzella, A., Napolitano, A., and d’Ischia, M. (2009) Efﬁcient synthesis of 5,6-dihydroxyindole dimers, key eumelanin building blocks, by a uniﬁed o-ethynylaniline-based strategy for the construction of 2-linked biindolyl scaffolds. J. Org. Chem., 74, 7191–7194. Okuda, H., Wakamatsu, K., Ito, S., and Sota, T. (2008) Possible oxidative polymerization mechanism of 5,6-dihydroxyindole from ab initio calculations. J. Phys. Chem. A, 112, 11213–11222.

References 35 d’Ischia, M., Napolitano, A., and Prota,

36

37

38

39

40

41

42

G. (1987) Sulfhydryl compounds in melanogenesis. Part I. Reaction of cysteine and glutathione with 5,6-dihydroxyindoles. Tetrahedron, 43, 5351–5356. Lambert, C., Chacon, J.N., Chedekel, M.R., Land, E.J., Riley, P.A., Thompson, A., and Truscott, T.G. (1989) A pulse radiolysis investigation of the oxidation of indolic melanin precursors: evidence for indolequinones and subsequent intermediates. Biochim. Biophys. Acta, 993, 12–20. Pezzella, A., Panzella, L., Crescenzi, O., Napolitano, A., Navaratman, S., Edge, R., Land, E.J., Barone, V., and d’Ischia, M. (2006) Short-lived quinonoid species from 5,6-dihydroxyindole dimers en route to eumelanin polymers: integrated chemical, pulse radiolytic, and quantum mechanical investigation. J. Am. Chem. Soc., 128, 15490–15498. Panzella, L., Pezzella, A., Napolitano, A., and d’Ischia, M. (2007) The ﬁrst 5,6-dihydroxyindole tetramer by oxidation of 5,5′,6,6′-tetrahydroxy- 2,4′-biindolyl and an unexpected issue of positional reactivity en route to eumelanin-related polymers. Org. Lett., 9, 1411–1414. Pezzella, A., Panzella, L., Natangelo, A.M., Arzillo, A.N., and d’Ischia, M. (2007) 5,6-Dihydroxyindole tetramers with “anomalous” interunit bonding patterns by oxidative coupling of 5,5′,6,6′-tetrahydroxy-2,7′-biindolyl: emerging complexities on the way toward an improved model of eumelanin buildup. J. Org. Chem., 72, 9225–9230. Arzillo, M., Pezzella, A., Crescenzi, O., Napolitano, A., Land, E.J., Barone, V., and d’Ischia, M. (2010) Cyclic structure motifs in 5,6-dihydroxyindole polymerization uncovered: biomimetic modular buildup of a unique ﬁvemembered macrocycle. Org. Lett., 12, 3250–3253. Kaxiras, E., Tsolakidis, A., Zonios, G., and Meng, S. (2006) Structural model of eumelanin. Phys. Rev. Lett., 97, 218102. Ito, S. and Nicol, J.A. (1974) Isolation of oligomers of 5,6-dihydroxyindole-2carboxylic acid from the eye of the catﬁsh. Biochem. J., 143, 207–217.

43 Memoli, S., Napolitano, A., d’Ischia, M.,

44

45

46

47

48

49

50

Misuraca, G., Palumbo, A., and Prota, G. (1997) Diffusible melanin-related metabolites are potent inhibitors of lipid peroxidation. Biochim. Biophys. Acta, 1346, 61–68. Jiang, S., Liu, X.M., Dai, X., Zhou, Q., Lei, T.C., Beermann, F., Wakamatsu, K., and Xu, S.Z. (2010) Regulation of DHICA-mediated antioxidation by dopachrome tautomerase: implication for skin photoprotection against UVA radiation. Free Radic. Biol. Med., 48, 1144–1151. Pezzella, A., Vogna, D., and Prota, G. (2002) Atropoisomeric melanin intermediates by oxidation of the melanogenic precursor 5,6-dihydroxyindole-2-carboxylic acid under biomimetic conditions. Tetrahedron, 58, 3681–3687. Pezzella, A., Vogna, D., and Prota, G. (2003) Synthesis of optically active tetrameric melanin intermediates by oxidation of the melanogenic precursor 5,6-dihydroxyindole-2-carboxylic acid under biomimetic conditions. Tetrahedron Asymmetry, 14, 1133–1140. Pezzella, A., Panzella, L., Crescenzi, O., Napolitano, A., Navaratnam, S., Edge, R., Land, E.J., Barone, V., and d’Ischia, M. (2009) Lack of visible chromophore development in the pulse radiolysis oxidation of 5,6-dihydroxyindole-2carboxylic acid oligomers: DFT investigation and implications for eumelanin absorption properties. J. Org. Chem., 74, 3727–3734. Napolitano, A., Crescenzi, O., and Prota, G. (1993) Copolymerization of 5,6-dihydroxyindole and 5,6-dihydroxyindole-2-carboxylic acid in melanogenesis: isolation of a crosscoupling product. Tetrahedron Lett., 34, 885–888. Okuda, H., Wakamatsu, K., Ito, S., and Sota, T. (2010) Regioselectivity on the cooxidation of 5,6-dihydroxyindole and its 2-carboxy derivative from the quantum chemical calculations. Chem. Phys. Lett., 490, 226–229. Zajac, G.W., Gallas, J.M., Cheng, J., Eisner, M., Moss, S.C., and AlvaradoSwaisgood, A.E. (1994) The fundamental

183

184

6 Structure of Melanins

51

52

53

54

55

56

57

58

59

unit of synthetic melanin: a veriﬁcation by tunneling microscopy of X-ray scattering results. Biochim. Biophys. Acta, 1199, 271–278. Cheng, J., Moss, S.C., Eisner, M., and Zschack, P. (1994) X-ray characterization of melanins – I. Pigment Cell Res., 7, 255–262. Cheng, J., Moss, S.C., and Eisner, M. (1994) X-ray characterization of melanins – II. Pigment Cell Res., 7, 263–273. Gallas, J.M., Zajac, G.W., Sarna, T., and Stotter, P.L. (2000) Structural differences in unbleached and mildly-bleached synthetic tyrosine-derived melanins identiﬁed by scanning probe microscopies. Pigment Cell Res., 13, 99–108. Clancy, C.M. and Simon, J.D. (2001) Ultrastructural organization of eumelanin from Sepia ofﬁcinalis measured by atomic force microscopy. Biochemistry, 40, 13353–13360. Pezzella, A., Napolitano, A., d’Ischia, M., Prota, G., Seraglia, R., and Traldi, P. (1997) Identiﬁcation of partially degraded oligomers of 5,6-dihydroxyindole-2carboxylic acid in Sepia melanin by matrix-assisted laser desorption/ ionization mass spectrometry. Rapid Commun. Mass Spectrom., 11, 368–372. Stark, K.B., Gallas, J.M., Zajac, G.W., Eisner, M., and Golab, J.T. (2003) Spectroscopic study and simulation from recent structural models for eumelanin: I. Monomers, dimers. J. Phys. Chem. B, 107, 3061–3067. Stark, K.B., Gallas, J.M., Zajac, G.W., Golab, J.T., Gidanian, S., McIntire, T., and Farmer, P.J. (2005) Effect of stacking and redox state on optical absorption spectra of melanins – comparison of theoretical and experimental results. J. Phys. Chem. B, 109, 1970–1977. Pezzella, A., Ambrogi, V., Arzillo, M., Napolitano, A., Carfagna, C., and d’Ischia, M. (2010) 5,6-Dihydroxyindole oxidation in phosphate buffer/polyvinyl alcohol: a new model system for studies of visible chromophore development in synthetic eumelanin polymers. Photochem. Photobiol., 86, 533–537. Pezzella, A., Iadonisi, A., Valerio, S., Panzella, L., Napolitano, A., Adinolﬁ, M.,

60

61

62

63

64

65

66

67

and d’Ischia, M. (2009) Disentangling eumelanin “black chromophore”: visible absorption changes as signatures of oxidation state- and aggregationdependent dynamic interactions in a model water-soluble 5,6-dihydroxyindole polymer. J. Am. Chem. Soc., 131, 15270–15275. Napolitano, A., Memoli, S., and Prota, G. (1999) A new insight in the biosynthesis of pheomelanin: characterization of a labile 1,4-benzothiazine intermediate. J. Org. Chem., 64, 3009–3011. Napolitano, A., Memoli, S., Crescenzi, O., and Prota, G. (1996) Oxidative polymerization of the pheomelanin precursor 5-hydroxy-1,4benzothiazinylalanine: a new hint to the pigment structure. J. Org. Chem., 61, 598–604. Costantini, C., Crescenzi, O., Prota, G., and Palumbo, A. (1990) New intermediates of phaeomelanogenesis in vitro beyond the 1,4-benzothiazine stage. Tetrahedron, 46, 6831–6838. Thomson, R.H. (1974) The pigments of reddish hair and feathers. Angew. Chem. Int. Ed. Engl., 13, 305–312. Napolitano, A., Di Donato, P., and Prota, G. (2001) Zinc-catalyzed oxidation of 5-S-cysteinyldopa to 2,2′-bi(2H-1,4benzothiazine): tracking the biosynthetic pathway of trichochromes, the characteristic pigments of red hair. J. Org. Chem., 66, 6958–6966. Land, E.J., Ito, S., Wakamatsu, K., and Riley, P.A. (2003) Rate constants for the ﬁrst two chemical steps of eumelanogenesis. Pigment Cell Res., 16, 487–493. Agrup, G., Hansson, C., Rorsman, H., and Rosengren, E. (1982) The effect of cysteine on oxidation of tyrosine, dopa, and cysteinyldopas. Arch. Dermatol. Res., 272, 103–115. Bush, W.D., Garguilo, J., Zucca, F.A., Albertini, A., Zecca, L., Edwards, G.S., Nemanich, R.J., and Simon, J.D. (2006) The surface oxidation potential of human neuromelanin reveals a spherical architecture with a pheomelanin core and a eumelanin surface. Proc. Natl. Acad. Sci. USA, 103, 14785–14789.

References 68 Peles, D.N., Hong, L., Hu, D.N., Ito, S.,

69

70

71

72

73

74

75

76

77

78

Nemanich, R.J., and Simon, J.D. (2009) Human iridal stroma melanosomes of varying pheomelanin content possess a common eumelanic outer surface. J. Phys. Chem. B, 113, 11346–11351. Palumbo, A., d’Ischia, M., Misuraca, G., Prota, G., and Schultz, T. (1988) Structural modiﬁcations in biosynthetic melanins induced by metal ions. Biochim. Biophys. Acta, 964, 193–199. Ito, S. and Fujita, K. (1985) Microanalysis of eumelanin and pheomelanin in hair and melanomas by chemical degradation and liquid chromatography. Anal. Biochem., 144, 527–536. Wakamatsu, K. and Ito, S. (2002) Advanced chemical methods in melanin determination. Pigment Cell Res., 15, 174–183. Ito, S. and Wakamatsu, K. (1998) Chemical degradation of melanins: application to identiﬁcation of dopaminemelanin. Pigment Cell Res., 11, 120–126. Napolitano, A., Vincensi, M.R., Di Donato, P., Monfrecola, G., and Prota, G. (2000) Microanalysis of melanins in mammalian hair by alkaline hydrogen peroxide degradation: identiﬁcation of a new structural marker of pheomelanins. J. Invest. Dermatol., 114, 1141–1147. Napolitano, A., Pezzella, A., d’Ischia, M., and Prota, G. (1996) New pyrrole acids by oxidative degradation of eumelanins with hydrogen peroxide. Further hints to the mechanism of pigment breakdown. Tetrahedron, 52, 8775–8780. Napolitano, A., Pezzella, A., Vincensi, M.R., and Prota, G. (1995) Oxidative degradation of melanins to pyrrole acids: a model study. Tetrahedron, 51, 5913–5920. Ward, W.C., Lamb, E.C., Cooden, D., Chen, X., Burinsky, D.J., and Simon, J.D. (2008) Quantiﬁcation of naturally occurring pyrrole acids in melanosomes. Photochem. Photobiol., 84, 700–705. Borovansky, J. and Elleder, M. (2003) Melanosome degradation: fact and ﬁction. Pigment Cell Res., 16, 280–286. Wakamatsu, K., Ito, S., and Rees, J.L. (2002) The usefulness of 4-amino-3hydroxyphenylalanine as a speciﬁc marker of pheomelanin. Pigment Cell Res., 3, 225–232.

79 Napolitano, A., Vincensi, M.R., d’Ischia,

80

81

82

83

84

85

86

87

M., and Prota, G. (1996) A new benzothiazole derivative by degradation of pheomelanins with alkaline hydrogen peroxide. Tetrahedron Lett., 37, 6799–6802. Fattorusso, E., Minale, L., Cimino, G., Stefano, S.D., and Nicolaous, R.A. (1969) Struttura e biogenesi delle feomelanine. Nota VI. Sulla struttura della gallofeomelanina-1. Gazz. Chim. Ital., 99, 29–45. Greco, G., Wakamatsu, K., Panzella, L., Ito, S., Napolitano, A., and d’Ischia, M. (2009) Isomeric cysteinyldopas provide a (photo)degradable bulk component and a robust structural element in red human hair pheomelanin. Pigment Cell Melanoma Res., 22, 319–327. Greco, G., Panzella, L., Napolitano, A., and d’Ischia, M. (2009) Biologically inspired one-pot access routes to 4-hydroxybenzothiazole amino acids, red hair-speciﬁc markers of UV susceptibility and skin cancer risk. Tetrahedron Lett., 50, 3095–3097. Costantini, C., Testa, G., Crescenzi, O., and d’Ischia, M. (1994) Photochemical ring contraction of dihydro-1,4benzothiazines. Tetrahedron Lett., 35, 3365–3366. Di Donato, P., Napolitano, A., and Prota, G. (2002) Metal ions as potential regulatory factors in the biosynthesis of red hair pigments: a new benzothiazole intermediate in the iron or copper assisted oxidation of 5-S-cysteinyldopa. Biochim. Biophys. Acta, 1571, 157–166. Ozeki, H., Ito, S., Wakamatsu, K., and Hirobe, T. (1995) Chemical characterization of hair melanins in various coat-color mutants of mice. J. Invest. Dermatol., 105, 361–366. Wakamatsu, K., Fujikawa, K., Zucca, F., Zecca, L., and Ito, S. (2003) The structure of neuromelanin as studied by chemical degradative methods. J. Neurochem., 86, 1015–1023. Lamoreux, M.L., Wakamatsu, K., and Ito, S. (2001) Interaction of major coat color gene functions in mice as studied by chemical analysis of eumelanin and pheomelanin. Pigment Cell Res., 14, 23–31.

185

wwwwwww

187

7 Properties and Functions of Ocular Melanins and Melanosomes Małgorzata Rózᠨanowska 7.1 Introduction

The color of human eyes is largely dependent on the amount, type, and distribution of melanin present in the iris [1, 2]. However, the iris is not the only part of the uveal tract that contains melanin (Figure 7.1). Melanin is in great abundance also in the remaining parts of the highly vascularized uvea – the choroid and the ciliary body. In the choroid, melanin is present in melanocytes, whereas in the iris and ciliary body melanin is present in two cell types: melanocytes in the stroma and in epithelial cells. Another melanin-containing epithelium in the eye is the retinal pigment epithelium (RPE), an essential monolayer of cells that separate the neurosensory retina from the choroidal blood supply. This chapter is focused on our current understanding of the properties and functions of melanin in the mammalian eye, particularly the human eye.

7.2 Biogenesis of Ocular Melanosomes and Melanogenesis

Ocular melanins, like most other natural melanins, are a mixture of brown/black eumelanin and red/brown pheomelanin. Melanin is synthesized in intracellular organelles, the melanosomes, which are present in uveal melanocytes or in specialized pigment epithelial cells of the uvea and retina. In the RPE the biogenesis of melanosomes and melanogenesis was followed and divided in several stages [4–6]. The ﬁrst stage involves budding off from the smooth endoplasmic reticulum (ER) of ovoid, membrane-bound organelles, called premelanosomes. Formation of premelanosomes occurs in the RPE early in fetal development, then ceases within a few weeks. Premelanosomes develop into stage I melanosomes, where the protein matrix is regimented, ﬁlaments stretch from one pole to the other, and the granule becomes more osmiophilic. Melanogenesis in the eye can occur only in the presence of functional tyrosinase [7], although a number of other proteins are likely to be involved such as tyrosinase-related protein (TRP)-1 and TRP-2 [8, Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

188

7 Properties and Functions of Ocular Melanins and Melanosomes

Figure 7.1 Diagram of a cross-section of the human eye depicting melanin-containing structures: RPE and the uveal tract composed of the choroid, ciliary body and iris (left). A photograph of the human fundus

showing the characteristic areas: macula and fovea with the highest density of photoreceptors responsible for acute vision (right). (Modiﬁed from [3].)

9], p locus protein [10], and lysosomal membrane surface proteins [11]. Tyrosinase encapsulated in coated vesicles is delivered to premelanosomes and stage I melanosomes. Melanogenesis in the eye starts very early during fetal development. Pigmentation in the RPE cells is present already in the fourth week of gestation. Polymerization of melanin within the RPE melanosomes continues until approximately 2 years of age in humans after which pigment turnover is believed to be quite low [5, 6]. Once full melanization has occurred, melanosome reaches the ﬁnal stage IV with no tyrosinase activity. Melanosomes in the melanocytes of the iris stroma in Caucasian eyes start developing pigmentation only after birth [4]. This can be observed easily in Caucasian babies born with blue irides, which may change color to brown within a few months after birth. Current understanding of pathways leading to biosynthesis of eumelanin and pheomelanin has been recently reviewed by Simon, Ito and colleagues [12, 13]. In short, both types of melanin derive from enzymatic oxidation of l-tyrosine catalyzed by tyrosinase to the common precursor dopaquinone (o-quinone of 3,4-dihydroxyphenylalanine (dopa)). In the absence of sulfhydryl compounds, dopaquinone undergoes intramolecular cyclization to form cyclodopa, which is then rapidly oxidized by a redox reaction with dopaquinone to give dopachrome (and dopa). Dopachrome then gradually rearranges to form 5,6-dihydroxyindole (DHI) and to a lesser extent DHI-2-carboxylic acid (DHICA). The DHI/DHICA ratio is determined by a dopachrome tautomerase (Dct; also known as TRP-2). Oxidation and subsequent polymerization of the dihydroxyindoles leads to the production of eumelanin. In the presence of cysteine, dopaquinone gives rise to cysteinyldopa isomers. Cysteinyldopas subsequently react with dopaquinone to

7.2 Biogenesis of Ocular Melanosomes and Melanogenesis

form cysteinyldopaquinones, eventually leading to the production of pheomelanin. The total amount of melanin produced is proportional to tyrosinase activity. It has been proposed that synthesis of natural melanins, which are a mixture of eumelanin and pheomelanin, proceeds in three distinct stages [12, 13]. In the ﬁrst stage, when the concentration of cysteine is above 0.13 μM, cysteinyldopa isomers are produced. When the concentration of cysteine is decreased but is still greater than 9 μM, cysteinyldopas are oxidized to produce pheomelanin. Only after most cysteinyldopas and cysteine are depleted is eumelanin produced in the third stage of melanogenesis. The ratio of eumelanin to pheomelanin is determined by tyrosinase activity, and the availability of tyrosine and cysteine in melanosomes. This theory of three-stage mixed melanogenesis is supported by studies on melanogenesis in cultures of human uveal and epidermal melanocytes [12, 13]. In these cells pheomelanin is produced at constant levels regardless of the degree of pigmentation, whereas eumelanin production is proportional to pigmentation. Another factor that affects melanin synthesis is melanosomal pH [12, 13]. Tyrosinase activity peaks at about pH 7.3 and is almost completely inactive under acidic conditions. It has been shown that melanosomes isolated from Caucasian skin exhibit acidic pH, while pH of melanosomes isolated from black skin is more neutral, and that difference corresponds to tyrosinase activities in these melanosomes and melanin production [14, 15]. By employing drugs to neutralize pH in melanosomes, the tyrosinase activity can be strongly upregulated and melanin content increased [14–16]. Considering that the cyclization of dopaquinone leading to eumelanin production proceeds slower at lower pH, while the cysteinyldopa– quinone cyclization leading to pheomelanin is faster, it has been suggested that neutral melanosomes will favor synthesis of eumelanin, while pheomelanin is more likely to be produced in acidic melanosomes [12]. Adult human RPE contains only mature melanosomes [6]. Consistently, adult bovine RPE cells exhibit very little (about 1/20 th of that in the choroid) activity of tyrosinase [17, 18]. In contrast, the adult bovine uvea exhibits considerable tyrosinase activity, particularly in the ciliary body where it is 2 times higher than in the choroid or iris [18]. Tyrosinase activity has been also detected in the ciliary body of the adult human eye [19]. The melanin content of the human choroid remains approximately constant with age, whereas melanin content in the human RPE decreases signiﬁcantly in aging eyes [20–23], supporting the view that melanin production is either absent in the adult human RPE or occurs only at a very slow rate. Interestingly, it has been demonstrated in the cultured RPE cell line ARPE-19 transfected with tyrosinase that melanogenesis can also occur in the absence of premelanosomes [24]. In the transfected cells, tyrosinase was associated with multivesicular and multilamellar organelles in which melanogenesis took place despite the fact that the organelles did not contain detectable amounts of proteins typical of premelanosomes, such as structural protein PMEL17 or TRP-1. These ﬁndings indicate that the classical pathway of melanogenesis, which occurs in four stages in melanosomes, is not essential for melanin synthesis and there is a possibility that melanin can be synthesized in the adult RPE via an alternative pathway also in vivo.

189

190

7 Properties and Functions of Ocular Melanins and Melanosomes

7.3 Melanin Content in Pigmented Structures of the Eye

The total melanin content as well as the relative content of pheomelanin to eumelanin varies in different melanin-containing parts of the eye. The following sections will discuss melanin composition in the RPE, choroid, and iris, and how it is affected by race and age. 7.3.1 Melanin Content in the RPE

RPE melanin is composed mostly of eumelanin [2]. Melanin content in the RPE in human eyes from 19 white and 16 black donors of different ages was studied by Weiter et al. by measurements of optical density of the RPE and their morphometric analysis [21]. To calculate optical density of the RPE, Weiter et al. measured intensities of 500- to 600-nm spectral light delivered from a rectangular aperture of 30 × 3 μm and transmitted through 8-μm thick sections of either the apical or the basal parts of the RPE. From these measurements the absorption coefﬁcients of the basal and apical portions of the RPE were calculated. The total optical density of the RPE monolayer was calculated as the product of the RPE height and the mean of the apical and basal absorption coefﬁcients. The optical densities were ascribed exclusively to melanin even though RPE cells contained lipofuscin likely to contribute to absorption and scatter of the incident 500- to 600-nm light. Nevertheless, the optical density measurements exhibited a positive statistical correlation with the counts of melanin granules. The measurements of the optical densities have demonstrated that the RPE exhibits a broad and ﬂat minimum in the paramacula, and increases almost 2-fold in both directions – towards the center of the macula and towards the equator, reaching in those locations values of about 0.3 ± 0.1 (Figure 7.2) [21, 25]. The highest melanin density in the RPE appears as a ring 1 mm proximal of the ora serrata, where the RPE terminates and is continuous with the ciliary body [5]. Morphological measurements have demonstrated that RPE melanin exhibits a highly polarized intracellular distribution in the RPE, particularly in young donors where most melanin is present in the apical portion of the cell [21]. It has been shown that there is no statistically signiﬁcant difference in melanin content in the RPE between whites and blacks [21]. In the RPE of both whites and blacks, melanin concentrations decrease with age [20, 21]. Feeney-Burns et al. studied the effect of donor age on RPE granule morphology and content in a 50 Caucasian cadaver eyes with ﬁve eyes per each decade of age [20]. The measurements demonstrated that there is a statistically signiﬁcant decrease with age in “pure” melanosomes accompanied by an increase of lipofuscin and complex granules: melanolysosomes and melanolipofuscin. The percentage area of the macular RPE cell occupied by melanosomes of about 8% in the ﬁrst two decades of life declines gradually to 3.5% of cell area in the 41–90 age group. The number of melanolipofuscin and melanolysosomes per RPE cell

7.3 Melanin Content in Pigmented Structures of the Eye a)

b)

Melanin (a) and lipofuscin (b) distribution in the RPE and choroid based on measurements of optical density (a) and ﬂuorescence (b). (Based on data from [21].)

Figure 7.2

section is considerable already in the ﬁrst decade of life (6.9 granules per cell section) and continues to rise to almost 15 complex granules per cell section in donors older than 61 years (Figure 7.3). As a percentage of cell area, the complex granules occupy 3.3% in the ﬁrst two decades and this rises to 8–10% of the RPE cell area in donors over 61 years old. In the 90- to 101-year-old maculas, most RPE melanin is present as melanolipofuscin [26]. At the equator, the effect of age on the percentage of RPE cell areas occupied by melanosomes and melanosome number is less pronounced than for the macular RPE (Figure 7.3) [20]. The area occupied by melanosomes decreases from 8% in the ﬁrst two decades to 5–6% in the elderly. Complex granules are rather sparse in the ﬁrst decade (0.8 granules per cell section). As a percentage of cell

191

192

7 Properties and Functions of Ocular Melanins and Melanosomes

Figure 7.3 Age-related changes in the content of pigment granules: melanosomes (Ms), melanolipofuscin (MLf), and lipofuscin (Lf) in human RPE. Mean number of

pigment granules per RPE cell section in the macular, equatorial and peripheral area of the retina from donors aged 1–20, 21–60, and 61–100 years. (Based on data from [20].)

area, the complex granules increase from 0.7% in the ﬁrst two decades to 4% in donors over 61 years old. In the farthest periphery (the last 4 mm before the ora serrata) the concentration of melanosomes is the highest in the youngest RPE with a mean of 34 granules per cell section (Figure 7.3) [20]. The percentage of RPE area occupied here by melanosomes decreases from 15% in the ﬁrst two decades to 4% in donors over 61 years old. The number of complex granules is the lowest here, but also signiﬁcantly increases with age. The percentage area occupied by melanolipofuscin and melanolysosomes increases from 0.5% in the 0- to 20-year group to 3.6% in the 61- to 100-year group. These morphological measurements of age-related decrease in melanin content in human RPE are overall consistent with results obtained by two other methods [22, 23]. Solubilization of RPE cells by sodium borohydride followed by centrifugation to separate the soluble part from the insoluble cellular debris provides a solution with the absorption spectrum identical to the absorption spectrum of synthetic eumelanin model [22]. As noted by the authors, the procedure leads to lipofuscin, melanolysosomes, and melanolipofuscin being pelleted during centrifugation, and, therefore, melanin present in the complex granules is excluded from the measurements. Based on absorption maximum at 260 nm of the solubilized fraction, melanin content has been calculated in human RPE from 61 eyes from donors ranging in age between 14 and 97 years. Fitting data to a linear regression shows that content of solubilized melanin decreases within eight decades of life about 3-fold. In another method of melanin quantiﬁcation, measurements of melanin free radicals by electron spin resonance (ESR) at pH 1 at liquid nitrogen temperature have been employed as a probe of melanin content in RPE cells scraped postmortem from the entire eye-cups of 105 donors [23]. The data obtained can be ﬁtted rea-

7.3 Melanin Content in Pigmented Structures of the Eye

sonably well using linear regression demonstrating that melanin content normalized to total cellular protein decreases 1.8-fold between the ages of 10 and 90 years. 7.3.2 Melanin Content in the Choroid

The optical density of choroid is maximal at the fovea and gradually decreases almost 2-fold towards the equator (Figure 7.2) [21]. The concentration of melanin in the choroid is on average 2.4 ± 1.0 times higher in the outer half of the choroid (adjacent to the sclera) than in the inner half (adjacent to the RPE). There is a highly signiﬁcant difference between whites and blacks in the optical density of the choroid [21]. The optical densities of the choroid for 500- to 600-nm light are 0.7 ± 0.5 and 1.4 ± 0.7 in the fovea, and 0.4 ± 0.3 and 0.7 ± 0.4 in the equator for whites and blacks, respectively. There is no statistically signiﬁcant change with age in the total choroidal optical density; however, there is a trend for the ratio of the outer/inner choroidal melanin density to increase with age [21]. These measurements of melanin density in postmortem RPE and choroid are consistent with measurements of optical density in vivo based on reﬂectance spectroscopy. From measurements of 675 nm light reﬂected from an area of 7 ° × 7 ° of the fundus incorporating the macula, the optical density of the RPE–choroid complex has been calculated as 0.79 ± 0.17 for 16 Caucasians, 1.36 for a black, and 0.06 for an albino [27]. Measurements of reﬂectance at 500 nm from smaller areas of the fundi in 10 subjects have shown that the optical densities of the RPE-choroid complex vary within the range of 0.79–8.5 in the fovea and 0.72–6.90 in the perifovea [28–30]. 7.3.3 Melanin Content in the Iris

The iris is the most anterior portion of the uveal tract and is the most visible. It is composed of three parts: an anterior limiting membrane, stroma, and the posterior pigment epithelium. As mentioned earlier, iridial melanosomes are present in stromal melanocytes and iris pigment epithelial (IPE) cells. Melanin from the IPE is similar in eyes of different color and it is mostly eumelanin [31]. The composition of iris stroma varies with respect to total melanin content and eumelanin/ pheomelanin ratio [31, 32]. Blue irides exhibit very small melanin content in the stroma, with no detectable eumelanin and pheomelanin content of only 0.03 μg/ iris. The stroma of green irides exhibits a total melanin content of 4.7 μg/iris and a eumelanin/pheomelanin ratio of 0.88. The stroma of green/blue mixed-color irides contains mostly eumelanin (17.8 μg/iris) and a very high eumelanin/ pheomelanin ratio of 44.5. Interestingly, the pheomelanin/eumelanin ratio of 3.7 in stromas of green/brown mixed-color and brown irides falls in between green and green/blue irides. The total melanin content is highest in stromas of brown irides (32.2 μg/iris).

193

194

7 Properties and Functions of Ocular Melanins and Melanosomes

While the color of the iris is largely determined by the variation in pigmentation in the iridial stroma, it is important to note that the iris color visible through the cornea results from different optical phenomena, including light absorption by different chromophores and multiple light scattering on melanosomes as well as other components of the connective tissue forming the stroma [2]. In albinism, where the melanin content in the pigment epithelium is markedly decreased or even absent, the color of irides may vary from yellow to a pink color. Studies of normal human donor eyes using light and electron microscopy have revealed that there is no signiﬁcant difference in the content and distribution of melanocytes in irides of different colors. The difference in iris color is mainly determined by the variation of the melanosome structure and distribution within the iridial melanocytes. Darker irides have larger melanin granules and greater granule density. Another study on quantiﬁcation of iridial melanin has been done on homogenates of whole human irides from donors of different age [33]. ESR measurements of melanin free radicals in the light blue irides showed no signiﬁcant age-related changes in the ESR signal parameters or melanin content. Considering that in these irides most melanin is expected to be localized to the IPE [31], it can be suggested that the content of IPE melanin does not change with age. Darkening of the iris can occur in the adult eye as a side effect of topical application of prostaglandin analogs, such as latanoprost, used to lower intraocular pressure in the treatment of glaucoma [34]. Morphometric studies have demonstrated that the only change induced by latanoprost occurs in the iris stroma where the existing melanosomes undergo a small enlargement and this is sufﬁcient to account for the darkening effect. Analysis of melanin composition of the iris stroma of cynomolgus monkeys subjected to 25–38 weeks of treatment with latanoprost has demonstrated that in the latanoprost-treated irides the amount of eumelanin increases 3- to 7-fold, while the pheomelanin content remains almost the same as in untreated irides [35]. As a result, latanoprost leads to the 3- to 5-fold increase in eumelanin/pheomelanin ratio in comparison with the control irides. Interestingly, latanoprost does not affect pigmentation of the iris epithelium.

7.4 Structure of Ocular Melanosomes

The structure of ocular melanosomes has been a subject of investigation for many years. The melanin composition of melanosomes from different ocular tissues has already been discussed. Early studies using transmission electron microscopy have shown that in ocular melanosomes, in similarity to other melanosomes, melanin is synthesized on a protein matrix and the whole granule is surrounded by a lipid bilayer. This section reviews the morphology of ocular melanosomes and their composition with regard to lipids and proteins.

7.4 Structure of Ocular Melanosomes

7.4.1 Morphology of Ocular Melanosomes

Melanosomes from different parts of the eye vary in shape, size, and age. Iridial stroma melanosomes in the young Rhesus monkey are cylindrical with the long axis often exceeding 2 μm and the short axis of about 0.2 μm [36]. With monkey aging there is progressive fusion of melanosomes forming aggregates composed initially of from two to four melanosomes and reaching numbers of 15–20 in the old monkeys; initially abundant long melanosomes seem to be replaced by shorter ovoid-shaped granules. Melanosomes from the human iridial stroma isolated from donors aged 30–67 years are rather uniform in size; small ovoid-shaped organelles with width and length dimensions of 0.25 ± 0.05 and 0.64 ± 0.10 μm, respectively [37]. In contrast, melanosomes from the human iris epithelium are spherical, with an average diameter of 1.02 μm [38]. Adult bovine melanosomes isolated from the whole iris–ciliary body complex seem to fall between these two with a long axis ranging from about 0.68 to 0.96 μm and a short axis ranging from 0.59 to 0.71 μm. Adult bovine melanosomes from the choroid also tend to have an ovoid shape with a long axis ranging from about 0.74 to 0.98 μm [39]. The choroidal melanosomes of the rat appear smaller and spherical with a diameter close to 0.4 μm [40]. Studies using transmission electron microscopy of age-related changes in choroidal melanosomes of Rhesus monkeys demonstrated that melanosomes in young animals are uniform in size and electron density, but with age there is an increasing number of melanosomes with altered morphology [41]. In aged monkeys almost all melanosomes appear as having an electron-dense “core” surrounded by material with very low electron density. No changes like these have been reported for humans. Choroidal melanosomes from human adults up to 67 years old appear similar to melanosomes from the human iridial stroma with width and length dimensions of 0.25 and 0.64 μm, respectively [37]. In the RPE of many species, including human, bovine, and rat, there are many elliptical or cigar-like melanosomes [20, 39, 40]. The long axis of the rat RPE melanosomes is close to 0.8 μm and the short axis is close to 0.4 μm [40]. Adult bovine RPE melanosomes tend to be larger than in the rat with a long axis within a range of about 0.6–2.9 μm and a short axis within a range of about 0.5–0.8 μm [39]. In the newborn bovine RPE there are considerably more cylindrical melanosomes than in the adult RPE. As discussed already, the most dramatic age-related changes in morphology of melanin-containing granules occur in the human RPE: the cylindrical melanosomes predominant in the newborn RPE seem to be disappear with aging so that in the adult RPE the melanosomes are more spherical and gradually seem to be replaced by complex granules – melanolysosomes and melanolipofuscin (Figure 7.3) [20, 26, 42]. Interestingly, in scanning electron microscopy, the morphology and size of the bovine ocular melanosomes appear unaffected by the procedure of chloroform– methanol extraction of melanosomal lipids [43]. It would be of interest to see the morphology of the aged human melanosomes, and melanolipofuscin/ melanolysosomes granules in particular, after such a procedure.

195

196

7 Properties and Functions of Ocular Melanins and Melanosomes

Imaging by atomic force microscopy of ocular melanosomes isolated from human and bovine iridial stroma and RPE reveals that these melanosomes, in a manner similar to the melanosomes isolated from the ink sac of the cuttleﬁsh Sepia ofﬁcinalis, or from human black and red hair, appear to be comprised of smaller substructures [44]. The substructures appear as nonporous spheres with lateral dimensions of a few tens of nanometers consistent with the structure of premelanosomes where zigzagging longitudinal strands form cross-links every 20 nm and melanin deposition occurs around these strands [45]. Additional information about melanosomal substructure has been elegantly deduced based on melanin composition, models of mixed melanogenesis, and comparisons of the surface photoionization thresholds of melanosomes isolated from dark brown and blue/green irides and from black and red hair using photoemission electron microscopy (PEEM) [44]. PEEM analysis of melanosomes isolated from black and red hair with a predominant content of eumelanin and pheomelanin, respectively, has demonstrated that they substantially differ in their ionization thresholds [44]. The ionization threshold for black hair eumelanosomes is 4.6 ± 0.2 eV (corresponding to an oxidation potential of −0.2 V versus normal hydrogen electrode (HNE)) [46, 47]. Red hair pheomelanosomes exhibit two ionization thresholds, one of which is the same as for eumelanosomes of 4.6 ± 0.2 eV and the other which is lower, 3.9 ± 0.2 eV (corresponding to an oxidation potential of +0.5 V versus HNE). Eumelanosomes of other origin, namely Sepia, bovine, and human RPE, also exhibit a single ionization threshold within the range of 4.4–4.8 (±0.2) eV. Melanosomes isolated from blue/green and dark brown irides are expected to have eumelanin/pheomelanin ratios of 1.3 and 14.8, respectively [32]. However, melanosomes from both dark brown and blue/green irides exhibited the same single ionization threshold of 4.9 ± 0.2 eV [44]. These results have been explained by a mixed melanogenesis model in which pheomelanin is initially synthesized and the pheomelanin core is covered by eumelanin exterior. Considering melanosomal substructure as a sphere of 30 nm diameter, the thickness of the eumelanin exterior would be ~9 and 3.6 nm for the dark brown and blue/green iridial melanosome, respectively. 7.4.2 Molecular Composition of Ocular Melanosomes

In addition to eumelanin and pheomelanin content, which has been discussed above, melanosomes also include a considerable fraction of proteins and lipids, which will be discussed below. 7.4.2.1 Melanosomal Proteins The proteomics of mature melanosomes is challenging because many melanosomal proteins are insoluble, presumably due to covalent binding to melanin [48]. Nevertheless, proteomic analyses have been applied to mature melanosomes and melanolipofuscin from porcine and human RPE, respectively, yielding a considerable number of proteins [48, 49]. To date, more than 100 proteins have been

7.4 Structure of Ocular Melanosomes

identiﬁed in RPE melanosomes. These include proteins involved in melanogenesis, organelle acidiﬁcation, organelle motility, cytoskeleton as well as proteolytic enzymes, channels, and transporters. Out of 102 proteins identiﬁed in the RPE melanosome by proteomics, only 12 proteins are common with the proteome of early stage melanosomes isolated from the human melanoma cell line MNT1. The majority of proteins in the RPE melanosome proteome are common with proteomes of other organelles, such as ER microsomes, mitochondria, Golgi apparatus, phagosomes, lysosomes, peroxisomes, the cytoskeleton, and the nucleus. Due to the small contamination of melanosomal preparations by mitochondria and other organelles, extramelanosomal origin of some of these proteins cannot be excluded. Unequivocal evidence for the presence of one of the lysosomal enzymes, cathepsin D, has been obtained by immunolabeling of cathepsin D in ﬁxed RPE and its detection by transmission electron microscopy. Cathepsin D was localized close to the surface of RPE melanosomes. As mentioned earlier, aging of the human RPE is accompanied by accumulation of complex granules: melanolysosomes and melanolipofuscin. Proteomics of human RPE melanolipofuscin has identiﬁed 110 proteins, 23 of which are common with the porcine RPE melanosomes and 18 with macrophage phagosomes [49]. Interestingly, only 14 proteins were identiﬁed as common for melanolipofuscin and lipofuscin studied by the same group. In another study of lipofuscin proteomics, 38 and 17 proteins common with human melanolipofuscin and porcine RPE melanosomes, respectively, have been identiﬁed [50]. Treatment of lipofuscin granules with sodium dodecyl sulfate (SDS) and/or proteinase K results in removal of all identiﬁable proteins and a 5-fold decrease in their amino acid content [50]. It would be interesting to see which proteins remain after the SDS/proteinase K treatment is applied to melanolipofuscin or melanosomes. 7.4.2.2 Melanosomal Lipids The melanin-rich center of the melanosome is surrounded by a lipid membrane. This membrane tends to disappear after melanosome isolation and puriﬁcation in a density gradient [48], imposing the requirement for a large number of melanosomes for lipid extraction. A thorough tandem mass spectrometry analysis of phospholipids was performed on lipids extracted from bovine ocular melanosomes isolated from three structures: iris–ciliary body complex, choroid, and RPE [43]. The analysis has revealed that the choroidal and iris–ciliary body melanosomes exhibit a similar composition of lipids with sphingomyelin, accounting for 54 and 41% of phospholipids in the choroid and iris, respectively. This is followed by glycerophosphocholine, lyso-glycerophosphocholine, lyso-glycerophosphoethanolamine, and phosphatidylglycerol. The fatty acid composition of these phospholipids is also rather similar in the choroid and iris. RPE melanosomes contain all of the lipids listed above; however, they exhibit striking differences in fatty acid composition in these phospholipids in comparison to the uveal melanosomes [43]. In the RPE, melanosomal phospholipids appear to contain more polyunsaturated fatty acyl chains than in the uvea. For example, in the RPE melanosomes, 93% of glycerophosphoethanolamine and 66%

197

198

7 Properties and Functions of Ocular Melanins and Melanosomes

of glycerophosphocholine contain long-chain polyunsaturated fatty acid species. In the iridial and choroidal melanosomes glycerophosphocholine contains at most 43 and 31% polyunsaturated lipids, respectively. The most unsaturated fatty acid of the mammalian body, docosahexaenoic acid, is esteriﬁed to 26.7% of glycerophosphoethanolamine and 9.6% of glycerophosphocholine in the RPE melanosomes. In contrast to uveal melanosomes, RPE melanosomes contain detectable quantities of glycerophosphoserine and glycerophosphate.

7.5 Role of Ocular Melanin as a Broadband Optical Filter

Absorption of light is considered as one of the most important functions of melanin in the eye [4, 6]. Melanins exhibit broadband absorption spectra increasing monotonically from the IR through visible to UV light. These spectra result from combined effects of light absorption and scatter. Direct measurements of single intact melanosomes with PEEM have allowed the determination of the product of the absorption coefﬁcient ε and the concentration of chromophores c absorbing 244 nm light [51]. For bovine RPE melanosomes isolated from a 1-week-old animal the product εc = 3895 ± 880 cm−1, whereas for melanosomes from an adult animal εc = 3515 ± 900 cm−1. The decrease of εc by a factor of 1.1 in the adult melanosomes compared to the newborn is consistent with an increase in the ratio of the eumelanin precursors DHI and DHICA. The [DHI]/[DHICA] ratio increases from 0.77 in newborn melanosomes to 1.25 in adult melanosomes. The absorption coefﬁcient of DHICA at 244 nm is about 3.8 times higher than that for DHI, so it can be calculated that, as a result of the increase of the [DHI]/[DHICA] ratio, the ratio of εc decreases by the same factor of about 1.1, provided the DHICA content and melanosome size remain the same. In addition to melanin, melanosomal proteins and their oxidation products are likely to contribute to the absorption of 244 nm light. Most of the light energy absorbed by melanin, particularly eumelanin, is dissipated as heat with only a tiny fraction released as ﬂuorescence, phosphorescence, or utilized for chemical changes such as formation of melanin free radicals and electron transfer to molecular oxygen [52]. 7.5.1 Role of the Iris as a Filter of Light

The human cornea absorbs UV-C and part of the UV-B radiation up to 295 nm [53], so only light above 295 nm reaches the iris and lens. The iris partly absorbs the remaining UV-B, UV-A, visible, and IR radiation so less light passes to the lens and the posterior segment comprised of the vitreous and retina. The iris regulates the size of the aperture through which light enters the more posterior parts of the eye. The iridial stroma is connected to the iris sphincter and dilator muscles. In response to light reaching the retina, either the sphincter or

7.5 Role of Ocular Melanin as a Broadband Optical Filter

the dilator contracts, in this was regulating the diameter of the circular opening in the center of iris – the pupil. The human pupil diameter depends on age and varies from 2–7 mm, when it is fully constricted in bright light, up to around 9 mm when it is fully dilated in the dark [54]. This pupillary light reﬂex plays an important role in adaptation of the eye to different levels of light, which normally vary within 11 orders of magnitude. By changing the size of the pupil, the pigmented iris can regulate the amount of light passing through the pupil up to 20-fold. While light enters the posterior segment of the eye mostly through the pupil, some light can be transmitted through the iris itself [55]. There are substantial differences in the transmittance through the iris itself depending on iris color [55]. In the rabbit, the average transmittance of freshly extracted blue irides is around 4.2 ± 1.7% and is greater by at least a factor of 4.2 than for brown irides. Blue irides transmit 7 times less light than albino irides. The difference in transmission properties of blue and brown irides seem to be less pronounced in humans where the ratio of transmittance for blue and brown irides isolated from human cadavers is only about 2.5 [55]. Transmission values of ﬂattened irides are quite considerable: 5.5 ± 2.8% for brown irides and 14 ± 6.3% for blue irides. Considering the surface of the iris and fully constricted pupil the amount of light passing through the iris in addition to that passing through the pupil can be estimated and compared for blue and brown irides. Calculation, based on a diameter of 11.7 mm of the opening through which light enters the iris [56] and a diameter of the constricted pupil of 2 mm [54], shows that the surface of the iris can be 34-fold greater than the pupil area. Considering the transmittances of human blue and brown irides cited above [55], it can be estimated that the light intensity entering the eye through the iris can be 1.9 and 4.8 times greater than the light entering the eye through the pupil for brown and blue irides, respectively. It should be noted that the human irides used for these measurements were used 2–3 days after enucleation and the IPE might have been dislodged due to extended storage [55]. Nevertheless, the study demonstrated that blue irides transmit 2.5–4.2 times more light than brown irides, and that this can account for a substantial part of light reaching the lens and posterior segment. These measurements of transmission of light through extracted irides are consistent with more recent psychophysical measurements of stray light in healthy human volunteers [57, 58]. The photoprotective role of iridial melanocytes is suggested by epidemiological studies of opacities in the lens known as cataract [59]. It has been shown that nuclear cataract and posterior subcapsular cataract are signiﬁcantly correlated with iris color, with higher a risk of cataract among medium brown, light brown, and yellow/green, in comparison with dark brown, irides. While there is no doubt pigmentation of the iris partly blocks light from reaching the posterior segment, it can also be suggested that absorption of light by iridial melanosomes can play a role in screening from light, and thus protection from oxidative and nitrative stress induced by UV and visible light, of different cell types present within the iris, including endothelial cells and erythrocytes as well as pigmented cells themselves [60–64]. Epidemiological studies of iridial melanoma support this protective function of melanosomes of the iridial stroma. Iridial

199

200

7 Properties and Functions of Ocular Melanins and Melanosomes

melanocytes are exposed to UV light above 295 nm transmitted through the cornea (Figure 7.1). Melanomas of the iris tend to originate from the inferior part of the iris where the exposure to sunlight is the greatest [2, 65]. The risk factors for the development of uveal melanoma include light iris color (blue, hazel, or gray), fair skin, inability to tan, and arc welding. It has been suggested that the high abundance of eumelanosomes in dark irides may act as an effective ﬁlter of light, thus providing protection from photo-oxidative stress and mutagenic effects. Light irides, on the other hand, do not exhibit a high density of stromal pigmentation. Moreover, blue/green and hazel irides contain a high proportion of pheomelanin to eumelanin, and therefore a thinner coating of eumelanin over the reactive pheomelanin core. It has also been suggested that exposure to sunlight may cause photodegradation of eumelanin and expose the more reactive pheomelanin, thus increasing the oxidative stress in the iridial melanocyte. 7.5.2 Role of Melanin in Light Transmission Through the RPE and Choroid

Light reaching the back of the adult human eye is mostly visible and IR covering the range from 390 to 1400 nm (Figure 7.1) [3]. UV light in the range 295–390 nm that passes through the cornea is almost completely absorbed by the lens. In young children there is a transmission window centered at about 320 nm with a transmission maximum of about 8%. As the lens ages, its transmission in the visible spectral range gradually decreases, with the effect being more pronounced the shorter the wavelength. Thus, the spectral characteristics of radiation incident on RPE and choroid are strongly dependent on the age and the presence of the crystalline lens. In the case of cataract surgery, the crystalline lens is replaced with an artiﬁcial lens, which could be either clear, transmitting all visible light above 390 nm, or yellow, mimicking the absorption of light by an old human lens. Melanosomes in the young RPE are localized in the apical portion of the RPE including its microvilli. The microvilli extend into the retinal layer containing photoreceptor outer segments with high concentrations of visual pigments. Here, in the apical portion of the RPE melanin can absorb light passing through photoreceptors thus minimizing light reﬂections and improving image quality. Based on optical densities measured by Weiter et al. of the RPE from cadaver eyes [21], it can be estimated that the RPE absorbs about 34–60% of incident light in the fovea, 21–40% in the paramacula, and 26–57% at the equator. As mentioned earlier, optical densities of choroidal melanin vary considerably between individuals, and there is a signiﬁcant difference between whites and blacks [21]. In the area underlying the fovea the choroid absorbs 37–94 and 80–99% of incident light in whites and blacks, respectively. At the equator, where the melanin density is the smallest, the choroid absorbs 21–80 and 50–92% of incident light in whites and blacks, respectively. Altogether, in the area of the fovea, the RPE and choroid absorb 58–97 and 87–100% of incident light for whites and blacks, respectively. At the equator, RPE

7.6 Antioxidant Properties of Ocular Melanin

and choroid absorb together 41–91 and 62–97% of incident light for whites and blacks, respectively. In addition to light entering the eye globe through the cornea, some light can be transmitted through the exposed part of the sclera adjacent to the cornea. Psychophysical measurements of volunteers with different degrees of ocular pigmentation have demonstrated that transmission of light through the anterior sclera–RPE–choroid complex is two orders of magnitude greater in people with blue irides than dark brown irides [58].

7.6 Antioxidant Properties of Ocular Melanin

Ocular melanin is present in structures exposed to light with high levels of oxygen and photosensitizers – an ideal environment for the production of reactive oxygen species (ROS). As discussed above, melanin can act as an efﬁcient natural sunscreen, thus protecting ocular tissues from adverse reactions induced by light. In addition, properties of synthetic melanin studied in suspension indicate that it can act as an efﬁcient scavenger of free radicals and quencher of electronically excited states of photosensitizers and singlet oxygen. Studies of both synthetic melanin and ocular melanosomes indicate that both can bind redox-active metal ions, such as iron or copper, and inhibit Fenton-type reactions catalyzed by these ions. These properties will be discussed below with regard to the potential functions of melanin in the eye. 7.6.1 Scavenging of Free Radicals

The ability of melanin to scavenge free radicals as well as reduce and oxidize other molecules is related to its redox properties. The key role in electron transfer is attributed to the functional subunits present in melanin: DHI, DHICA, and their fully oxidized (quinone) and semioxidized (semiquinone) forms in eumelanin, and o-aminophenols, such as 1,4-benzothiazine and their corresponding fully oxidized o-quinonimine and semioxidized o-semiquinonimine in pheomelanin [52, 66]. However, these subunits present in the melanin polymer exhibit much smaller reactivity in comparison to the very reactive free o-quinones and o-semiquinones. The relatively small reactivity of melanin subunits may be ascribed to their restricted accessibility and/or modiﬁed ionization potentials and electronic afﬁnity. From pulse radiolysis measurements of interaction of synthetic melanins with a number of oxidizing and reducing radicals it appears that the synthetic model of eumelanin requires stronger reducing radicals than the synthetic model of pheomelanin [67]. The one-electron reduction potential of the eumelanin model is about 0.52 V versus NHE; whereas for the pheomelanin model, the reduction potential is close to 0.35 V versus NHE. Assuming that all monomer units of the

201

202

7 Properties and Functions of Ocular Melanins and Melanosomes

eumelanin are available for redox reactions, it can be estimated that about 25% of the eumelanin sites are available for reduction, while 75% of the sites are available for oxidation. Synthetic models of eumelanin have also been investigated by electrochemistry yielding potentials of +460 and +525 mV versus saturated calomel electrode (SCE) for a two-electron oxidation and +20 and −355 mV versus SCE for a two-electron reduction [68]. Synthetic melanins can scavenge effectively a number of free radicals that can be formed in the eye, such as the hydroxyl radical, superoxide radical, nitrogen dioxide radical, or carotenoid cation radical [67, 69]. Hydroxyl radicals can be formed as a result of decomposition of hydrogen peroxide by redox active metal ions such as iron or copper [70]. In the cellular environment there is a high abundance of molecules with which •OH can react with bimolecular rate constants approaching the diffusion-controlled limits. Due to this extremely high reactivity of the hydroxyl radical in the cell, the ability of melanin to scavenge •OH is likely to play a role only if •OH is generated within the melanosomes or close proximity. Superoxide radicals are formed in all cells as a byproduct of mitochondrial respiration. In addition, in the RPE, superoxide is formed during phagocytosis of photoreceptor outer segments, or by photoexcited melanosomes or lipofuscin [71, 72]. Oxidation of melanin-bound metal ions such as Fe(II) or Cu(I) by molecular oxygen also leads to generation of superoxide [66]. Interaction of melanin with the superoxide radical leads either to its reduction to hydrogen peroxide or oxidation to molecular oxygen. Thus melanin can be viewed as a pseudo-dismutase. NO2ᠨ is formed as a result of interaction of long-lived nitric oxide with oxygen or as a result of oxidation of nitrite [73]. Nitric oxide, which is generated both in the retina and choroid, has a relatively long lifetime in tissues and therefore it may diffuse near melanosomes. It can be suggested that in proximity to melanosomes it may encounter superoxide, forming another reactive nitrogen species (RNS), peroxynitrite. Even though peroxynitrite is not a free radical, it is highly reactive with proteins and lipids, leading to their nitration. Scavenging of peroxynitrite by a synthetic model of neuromelanin has been demonstrated [74, 75]. Considering that protein nitration occurs in the retina and is substantially increased as a result of exposure to light or in retinal diseases [76], it is of high importance to investigate interactions of ocular melanins and melanosomes with RNS. The ability of melanin to scavenge the carotenoid radical cation may be important in human RPE where lutein, zeaxanthin, and β-carotene are present [77]. These carotenoids are believed to play a protective role in the RPE; however, they may become pro-oxidant and cytotoxic upon oxidation. Semioxidized forms of all these carotenoids are scavenged by synthetic models of eumelanin and pheomelanin [69], presumably by electron/hydrogen transfer, thus regenerating the parent compound. It remains to be shown whether or not RPE melanosomes can also protect from carotenoid degradation. An important question is whether melanins can scavenge peroxyl radicals propagating lipid peroxidation. While the interaction of melanins with lipid-derived

7.6 Antioxidant Properties of Ocular Melanin

peroxyl radicals has not been studied directly, the efﬁcient scavenging of peroxyl radicals derived from methanol by synthetic models of eumelanin and pheomelanin, suggests this possibility [78]. As mentioned earlier, RPE melanosomes contain polyunsaturated lipids. Oxidation of these lipids can lead to the formation of a number of noxious species, including some small polar aldehydes that can diffuse outside the granule. Thus, the ability of melanin to inhibit lipid peroxidation within RPE melanosomes would be a highly desirable function. While melanin structure appears very complex and resistant to elucidation, the structure of melanosomes is even more complex. As already mentioned, redox properties of whole melanosomes have been studied using PEEM [44]. PEEM is inherently a surface technique with electrons originating from no further than within a few nanometers of the melanosomal surface. This can be viewed as an advantage as interaction of melanosomes with many extragranular molecules is likely to proceed at the surface. As mentioned earlier, PEEM results have revealed that all ocular melanosomes studied so far, bovine and human RPE melanosomes, and human iridial stroma melanosomes isolated from dark brown and blue/green irides, exhibit similar single photoionization thresholds ranging from 4.4 ± 0.2 to 4.9 ± 0.2 eV (corresponding to 253–282 nm wavelength of light and oxidation potential of about −0.2 V versus NHE). This single photoionization threshold seems to be typical for eumelanosomes. Red hair pheomelanosomes exhibit an additional photoionization threshold of 3.8 ± 0.2 eV corresponding to 326 nm wavelength of light and oxidation potential of +0.5 V versus NHE) [44, 46, 47]. 7.6.2 Quenching of Electronically Excited States of Photosensitizers and Singlet Oxygen

Interaction of synthetic melanins with electronically excited states of photosensitizers or with the excited state of molecular oxygen, known as singlet oxygen (1Δg), can lead to transfer of the excess energy to melanin [66]. This effect suggests that melanin can offer photoprotection not only as a passive absorber of light, but also by preventing photosensitized oxidation. The energy transfer from electronically excited states is particularly efﬁcient for positively charged photosensitizers, such as cationic derivatives of porphyrins, which bind to melanins. When a melanin-bound photosensitizer absorbs light, the energy transfer occurs within femtoseconds from the excited singlet state of the photosensitizer to melanin [79]. As a result, the photosensitizer molecule returns to the ground state, and is unable to undergo intersystem crossing and form the excited triplet state – the usual culprit responsible for subsequent interactions with oxygen and formation of damaging ROS [3]. It has been demonstrated that human RPE melanosomes exhibit similar ability to synthetic melanins to bind cationic photosensitizers and strongly inhibit their photosensitizing properties (Figure 7.4) [80, 81]. Quenching of excited states of photosensitizers by melanosomes may be important in vivo in the presence of photosensitizing molecules of both exogenous and

203

204

7 Properties and Functions of Ocular Melanins and Melanosomes

Effect of native human RPE melanosomes isolated from donors 60–90 years old (HMs) and photodegraded human RPE melanosomes (dHMs) on photosensitized oxidation of histidine mediated by cationic derivative of porphyrin. Samples

Figure 7.4

contained melanosomes at indicated concentrations, 88 μM porphyrin, 2 mM histidine, and 0.1 mM ESR oximetry spin probe. Human RPE melanosomes were photodegraded by exposure to 5.54 kJ/cm2 of blue light (390–490 nm). (Modiﬁed from [80].)

endogenous origin. Xenobiotics with photosensitizing properties and high afﬁnity for binding to melanin include chloroquinone derivatives used as antimalarial drugs [82–84]. These antimalarial drugs cause phototoxic effects in the skin and the eye, and the ocular effects of their binding to melanin and accumulation in the eye are discussed below. Other drugs that deserve consideration with respect to interactions with melanin are drugs used for phototherapy where their photosensitizing properties are highly desired. When phototherapeutic drugs are targeted to affect melanin-free tissues, their binding to melanin and quenching of their excited states may be seen as protective for pigmented cells. For example, photodynamic therapy of choroidal neovascularization is meant to destroy pathological blood vessels growing into the retina in the course of the “wet” form of age-related macular degeneration (AMD). However, the photosensitizing drug used for that therapy, verteporﬁn (Visudyne®) tends to enter the RPE causing, in some cases, particularly after multiple treatments, collateral damage to this extremely important monolayer of postmitotic cells essential for vision [85]. It would be of interest to investigate the possibility that RPE melanosomes/melanolipofuscin bind to verteporﬁn (or other newly developed photosensitizers) and moderate their photosensitizing action. It could be suggested that during initial treatments, this binding could indeed play a protective role. However, if the photosensitizer remains bound to melanin, the

7.6 Antioxidant Properties of Ocular Melanin

binding capacity of melanin could be exceeded during subsequent treatments. Moreover, while binding of photosensitizers to melanosomes may be protective, it is unknown what the effects are of binding of photosensitizers to melanolipofuscin. Understanding the interactions between melanin-containing granules and photosensitizing drugs could allow better design of photodynamic treatment. Melanolipofuscin is expected to contain similar photosensitizer(s) as those present in lipofuscin granules [86]. It can be suggested that due to the proximity of melanin to these photosensitizers in melanolipofuscin granules, melanin can quench their photoexcited states, thus reducing the photoreactivity of melanolipofuscin. Whether or not this energy transfer occurs in melanolipofuscin still awaits experimental conﬁrmation. Another molecule present in lipofuscin is a pyridinium bisretinoid (so-called A2E), which, because of its positive charge, can be expected to bind to melanin. Although A2E provides an almost negligible contribution to photosensitizing properties of whole lipofuscin [3, 86], quenching by melanin of its excited state may be beneﬁcial by preventing A2E photodegradation that leads to formation of deleterious degradation products. Determination of whether or not photodegradation of A2E is slower in melanolipofuscin than in lipofuscin also needs experimental testing. Interestingly, it has been demonstrated that the inhibitory effect of human RPE melanosomes on photosensitized oxidation is substantially enhanced in the case of photodegraded melanosomes (Figure 7.4) [80]. Photodegradation was induced by a dose of blue light corresponding to 4.4 years of daily exposure of the human eye to ambient light. Under conditions where nondegraded melanosomes inhibited photosensitized oxidation of histidine mediated by cationic porphyrin by about 42%, the inhibition in suspension of degraded melanosomes was greater than 90%. It needs to be stressed that in this experiment melanosome photodegradation induced by blue light was rather extensive leading to a 2.3-fold decrease in the intrinsic free radical signal of melanin. Thus, it is intriguing why, despite the substantial degradation of melanin polymer, melanosome ability to inhibit photosensitized oxidation is enhanced. One possibility, which requires further investigation, is that access to melanin binding sites becomes facilitated upon melanosome photodegradation. It has been demonstrated that synthetic melanins and RPE melanosomes solubilized at pH 12 (and thus most likely oxidized) can quench the excited state of molecular oxygen that can be formed as a result of energy transfer from photoexcited photosensitizers to molecular oxygen [3, 66, 87]. In contrast to ground state oxygen, singlet oxygen can readily oxidize proteins, nucleic acids and unsaturated lipids. Yet, the cellular defenses against singlet oxygen seem to be rather limited. In contrast to other ROS, such as superoxide radical anion, hydrogen peroxide, or lipid hydroperoxides, no enzymes have evolved to deactivate singlet oxygen. Thus, melanin as a singlet oxygen quencher is potentially important for all pigmented tissues exposed to light. However, it can be suggested that due to the relatively short lifetime of singlet oxygen, which in water is only about 3–4 μs, and the abundance of substrates of oxidation in the cellular environment, the quenching

205

206

7 Properties and Functions of Ocular Melanins and Melanosomes

of singlet oxygen by melanin is likely to occur only if singlet oxygen is generated in proximity or within melanin-containing granules. These conditions are probably met in melanolipofuscin; however, experimental proof is still lacking. 7.6.3 Sequestration of Redox-Active Metal Ions

Melanin has the capacity to bind and accumulate many metal ions including redoxactive metals such as iron and copper [4, 52, 88]. Iron and copper can be damaging by catalyzing the decomposition of hydrogen peroxide and lipid hydroperoxide that result in the formation of the highly reactive free hydroxyl radical and/or initiation of a chain of lipid peroxidation [70]. It has been demonstrated that binding of iron and copper ions to melanin diminishes their ability to catalyze the generation of free hydroxyl radicals and lipid peroxidation [4, 52]. Even though melanin complexes with Fe(II) ions are readily oxidized by hydrogen peroxide and oxygen, the hydroxyl radicals produced are mostly scavenged by melanin and few leak out of the melanin polymer. Once iron ions are oxidized, melanin-bound Fe(III) becomes substantially more difﬁcult to reduce by mild biological reductants such as NAD(P)H or ascorbate in comparison with free Fe(III) ions or Fe(III) ions bound to ADP or citrate. Thus, melanin can prevent redox cycling of iron ions and their action as catalysts in Fenton-type reactions. It has been shown that ocular melanosomes isolated from human, porcine, and bovine RPE can also bind iron ions, and exhibit similar protective effects as synthetic melanins [80, 81, 89, 90]. A very convincing example of protective action of melanin against iron-catalyzed oxidation has been obtained by comparison of ironcatalyzed oxidation in a suspension of bovine RPE homogenate isolated from pigmented and nonpigmented parts of bovine fundus (Figure 7.5) [80]. To test whether or not the presence of melanin in pigmented cells was responsible for the inhibitory effect, the nonpigmented RPE cells were supplemented with either bovine melanosomes or synthetic eumelanin. As a result of addition of bovine melanosomes, the rate of oxidation in suspension of nonpigmented cells was slowed down to a similar level as in pigmented cells. Human RPE melanosomes isolated from 60- to 90-year-old donors are also effective in inhibiting iron-catalyzed oxidation (Figure 7.5). These ﬁndings are of particular importance considering that RPE cells provide the blood–retina barrier, and therefore are heavily involved in continual trafﬁcking of iron between the choroidal blood supply and photoreceptors [91, 92]. Iron delivered from blood to the RPE is partly used there as a cofactor for mitochondrial and other enzymes, such as enzymes essential for the regeneration of visual pigments. A portion of iron is transported through the RPE to photoreceptors and incorporated in their outer segments. The distal tips of photoreceptor outer segments are phagocytosed daily by the RPE, providing another source of iron. Due to their pro-oxidant properties, the free iron ion concentrations in the cell are meant to be maintained at very low levels by efﬁcient homeostatic mechanisms,

7.6 Antioxidant Properties of Ocular Melanin a)

Effect of bovine (a) and human (b) RPE melanosomes on oxidation catalyzed by iron ions. Oxidation was induced by Fe-ADP/ascorbate (0.05 mM/0.20 mM) in a suspension of bovine RPE homogenate isolated from pigmented (BP) and nonpigmented (BNP) parts of bovine fundus (a). To test whether the presence of melanin in pigmented bovine fundus was responsible for the inhibitory effect, either bovine

Figure 7.5

b)

melanosomes (BMs) or synthetic eumelanin (DM) were added to nonpigmented bovine RPE at concentrations corresponding to concentration of melanin of 30 or 60 μg/ml. Inhibitory effect of human RPE melanosomes (HMs) was tested on iron ion-catalyzed lipid oxidation in a suspension of 10 mM linoleate (LA), 0.2 mM ascorbate, and 0.05 mM Fe-ADP (b). (Modiﬁed from [80].)

controlling its compartmentalization and regulating its release and trafﬁcking. However, it can be suggested that due to its physiological functions RPE is at continual risk of exposure to iron ions that escape from iron-binding proteins. With aging there is a substantial increase in iron content in the RPE and choroid [93]. Iron content is further increased in several retinal degenerations. For example, there is a 5-fold increased iron in the RPE affected by AMD than in age-matched healthy RPE [94]. Another factor likely to contribute to increasing the risk of Fenton-type processes in the aged RPE is age-related decrease of activity of catalase – the major enzyme responsible for decomposition of hydrogen peroxide [95]. It remains to be investigated in which intracellular compartment(s) of the RPE the accumulation of iron occurs and whether melanosomes can protect RPE cells from deleterious effects of iron. 7.6.4 Testing Protective Effects of Ocular Melanin in Cultured Cells

All the antioxidant properties discussed above indicate that melanosomes may play a protective role in the eye in vivo. However, providing experimental evidence on animals with different degrees of ocular pigmentation to support this notion has

207

208

7 Properties and Functions of Ocular Melanins and Melanosomes

proven to be unexpectedly difﬁcult and conﬂicting results have been obtained [96]. Also, attempts to demonstrate the protective effect of RPE melanosomes in cultured cells have proven so far unsuccessful [97, 98]. Susceptibility to apoptosis of primary cultures of fetal human RPE cells with different degrees of pigmentation has been studied by Seagle et al. [97]. Melanin content was assessed by spectrophotometry. The intensity of melanin free radicals in the dark and upon exposure to increasing intensity of 355-nm laser light was monitored by ESR. To assess potential photoprotective effects of melanin, cells were exposed for 7 days to continuous blue light irradiation (440 nm, 4.55 mW/ cm2) while cells kept in the dark served as controls. Staining of exposed phosphatidylserine with Annexin V was used as a measure of apoptosis. It has been stated in the results that the RPE cells less susceptible to apoptosis have relatively low optical absorption spectra even though the intensities of the ESR signals of melanin are relatively high [97]. Plotting the data points shows that the cell cultures employed exhibited considerable variability and linear regression analysis of the data provide low correlation coefﬁcients (Figure 7.6). It can be argued that the higher amplitudes of melanin free radicals measured for some RPE cells could reﬂect a lower content of melanin-bound iron and/or higher concentrations of reductants such as ascorbate [98, 99], each of which could also confer protection from light-induced apoptosis. In conclusion, the role of endogenous melanosomes in protection of the RPE against light damage requires further investigation. Zareba et al. have undertaken a thorough investigation of effects of exogenous melanosomes on the human RPE cell line, ARPE-19 [98]. The cells were fed with bovine or porcine melanosomes to accumulate intracellular granule concentrations of more than 20 melanosomes/cell and then exposed to visible light or oxidants. Several endpoints were evaluated including integrity of the plasma membrane, mitochondrial activity, cell adhesion, and lysosomal integrity, and compared with cells fed with melanosomes, charcoal, silica particles, and control cells. The results have demonstrated that the presence of intracellular melanosomes neither prevented nor exacerbated the cytotoxic effects of hydrogen peroxide, tert-butyl hydroperoxide, and visible light. However, it can be argued that the lack of effect of melanosomes could be due to the majority of oxidants being formed in the extracellular culture medium. The culture medium contained iron ions, and therefore it is likely that the decomposition of tert-butyl hydroperoxide and hydrogen peroxide occurred mainly extracellularly. The riboﬂavin present in the culture medium used could also be a source of extracellular phototoxic effects as observed in cultured cells exposed to UV-A or blue light [100]. Furthermore, phagocytosed melanosomes may be surrounded by an additional lipid membrane thereby hindering their interaction with ROS. Under physiological conditions RPE cells are likely to be exposed to oxidative stress due to accumulation of lipofuscin [3, 86]. It would be of interest to test the effects of melanosomes on oxidative stress induced inside the RPE cell by photoexcited lipofuscin.

7.7 Pro-Oxidant Properties of Ocular Melanosomes a)

b)

d)

Susceptibility to apoptosis in primary cultures of fetal human RPE cells after 7 days of exposure to blue light (circles) or kept in the dark (triangles), and arithmetical differences between apoptosis values of cells exposed to light and kept in the dark (rectangles) as functions of optical density of RPE cells measured at 500 nm (a), amplitude of melanin radical in the dark (b) or melanin

Figure 7.6

c)

e)

radical photoinduced by exposure to laser light (250 mW; 355 nm) (c). (d and e) Amplitudes of melanin radical in the dark (d) or during exposure to laser light (e) as a function of optical density of RPE cells. Linear regression lines and their corresponding correlation coefﬁcients are included in diagrams. (Based on data from [97].)

7.7 Pro-Oxidant Properties of Ocular Melanosomes

Ocular melanosomes exhibit several pro-oxidant properties. They can be a source of ROS, induce depletion of important cellular reductants, and facilitate Fentontype reactions. This can be another reason why it is so difﬁcult to observe protective effects of melanosomes in cultured cells or in vivo, where the antioxidant action can be counterbalanced by the pro-oxidant action of melanosomes. Next, the prooxidant properties of ocular melanosomes and their age-related changes will be discussed.

209

210

7 Properties and Functions of Ocular Melanins and Melanosomes

7.7.1 Generation of ROS and Oxidation of Cellular Reductants

Photoexcitation with UV or visible light of ocular melanosomes results in similar effects as those observed for synthetic melanins: generation of photoinduced melanin free radicals and formation of ROS, such as superoxide radical and hydrogen peroxide [66, 71]. While for synthetic eumelanin most oxygen consumed during irradiation with visible light is converted to hydrogen peroxide, only around 35% of consumed oxygen is converted to hydrogen peroxide for RPE melanosomes isolated from donors younger than 33 years. The ratio of hydrogen peroxide produced to total oxygen consumed decreases with donor age. For donors older than 78 only 13% of oxygen consumed during irradiation of RPE melanosomes accumulates as hydrogen peroxide. This indicates that RPE melanosomes are susceptible to photo-oxidation of intragranular components. Consistently, upon irradiation of human RPE melanosomes with visible light accumulation of lipid hydroperoxides can be detected. Light-induced oxidation is strongly dependent on the irradiation wavelength. The rate of photoinduced oxygen uptake in suspension of human or bovine RPE melanosomes increases with decreasing wavelength measured in the range of 300 to 600 nm (Figure 7.7) [101]. While for UV and short-wavelength blue light the rate of photo-oxidation of human RPE melanosomes is slower than for lipofuscin by almost an order of magnitude, the difference becomes smaller at longer wave-

Figure 7.7 Wavelength dependence of susceptibility of bovine (BMs) and human (HMs) RPE melanosomes to photo-oxidation and their comparison with human RPE lipofuscin (HLf). Human granules were

isolated from donors above 60 years old. The initial rates of oxygen uptake have been normalized to equal number of incident photons. (Modiﬁed from [102].)

7.7 Pro-Oxidant Properties of Ocular Melanosomes a)

c)

b)

Figure 7.8 Photogeneration of superoxide by human RPE melanosomes isolated from donors less than 40 (MS < 40) or more than 80 years old (MS > 80). ESR spectra recorded in samples containing melanosomes (0.9 × 109 granules/ml) and 0.2 M spin trap 5,5-dimethylpyrroline-N-oxide (DMPO) in dimethyl sulfoxide before (a, b), and after (d, e) 22 min of irradiation with broadband blue light. (c) Kinetics of changes

d)

e)

in amplitude of DMPO-OOH spin adduct formed during irradiation. Arrow: beginning of irradiation. Incubation of melanosomes with DMPO in the dark did not result in the appearance of an ESR signal of spin adducts. Instrument settings: time constant 328 ms; sweep time 160 s; microwave power 10 mW; modulation amplitude 1.0 G. (Modiﬁed from [71].)

lengths. At wavelengths longer than 550 nm human RPE melanosomes are more photoreactive than lipofuscin. Interestingly, the susceptibility to photo-oxidation and the light-induced generation of superoxide radical by human RPE melanosomes substantially increases with donor age (Figure 7.8) [71]. Irradiation with blue light of suspension of the RPE melanosomes from donors more than 80 years old induces 2.4-fold faster oxygen uptake than for melanosomes from donors less than 40 years old. Despite the 40% increase in production of superoxide by older melanosomes in comparison to young melanosomes, there is no signiﬁcant difference in accumulation of the product of superoxide dismutation, hydrogen peroxide. This indicates that with age melanosomes may lose their ability to reduce superoxide and/or gain an ability to decompose H2O2. It has been shown that photoexcited human RPE melanosomes can oxidize extragranular unsaturated lipids (Figure 7.9) [71, 103]. Generation of hydrogen peroxide by photoexcited melanosomes from the RPE or iris strongly increases in the presence of physiological reductants, such as ascorbate or NADH, whereas the reductants are depleted (Figures 7.9 and 7.10) [71, 99, 104–107]. Altogether, the properties of isolated melanosomes indicate that they may contribute to oxidative stress in the RPE by generation of reactive species and depleting cellular reductants.

211

212

7 Properties and Functions of Ocular Melanins and Melanosomes

Figure 7.9 The effect of the addition of liposomes with polyunsatuarated lipids (PUFA), bovine serum albumin (BSA), NADH, or mM ascorbate (AscH) on the rate of broadband, blue-light-induced oxygen uptake in suspensions of human RPE melanosomes (MS) or lipofuscin (LF).

Control samples were with melanosomes or lipofuscin without additional substrate of oxidation (CTRL). The rates of oxygen uptake in the dark were negligible for all samples studied. Signiﬁcant differences are relative to the controls. *P < 0.05, **P < 0.01. (Modiﬁed from [71].)

a)

b)

Depletion of ascorbate (a) and formation of hydrogen peroxide (b) during aerobic irradiation with blue light of bovine RPE melanosomes (BMs) in the presence of ascorbate (Asc). Controls show levels of ascorbate during incubation with bovine RPE

melanosomes in the dark (a) and formation of hydrogen peroxide during incubation of bovine RPE melanosomes with ascorbate in the dark and during irradiation of bovine RPE melanosomes in the absence of ascorbate (b). (Modiﬁed from [99].)

Figure 7.10

7.7 Pro-Oxidant Properties of Ocular Melanosomes

7.7.2 Pro-Oxidant Effects of Interactions of Melanosomes with Metal Ions

While melanin can effectively inhibit redox-cycling of metal ions by cellular reductants and decomposition of hydrogen peroxide and lipid hydroperoxides catalyzed by metal ions, melanin itself can reduce Fe(III) or Cu(II) ions when they are complexed with a strong chelator and/or when they exceed the number of melanin binding sites [108]. It remains to be investigated how close to saturation with iron melanosomes in the aged RPE are, especially if affected by AMD. Superoxide radical, which can be generated by photoexcited melanin/ melanosomes, can also reduce iron and copper ions. Oxidation of melanin-bound Fe(II) or Cu(I) by hydrogen peroxide or molecular oxygen leads to the formation of hydroxyl radical and superoxide radical, respectively. As both radicals are generated within the melanin granule, they are likely to be scavenged by melanin. However, in cells such as RPE, life-long exposure to visible light and scavenging of radicals may lead to melanin degradation. Degradation of isolated RPE melanosomes can be easily achieved by their exposure to visible light [80, 89, 90]. For human RPE melanosomes isolated from donors 60–90 years old, the dose of blue light leading to 2.3-fold decrease in melanin free radical was equivalent to 4.4 years of daily exposures to ambient light [80]. It is still an open question whether melanosome degradation occurs in vivo or whether cellular antioxidants are able to prevent or minimize it. With aging, RPE melanosomes undergo substantial changes with regard to their morphology, melanin content, and photophysical and photochemical properties. Previous studies have shown that photodegradation of melanosomes mimics several changes observed in the aging of human RPE melanosomes, including (i) an increase in ﬂuorescence [23, 109], (ii) a decrease in intrinsic melanin free radicals [80, 90, 110], and (iii) an increase in photoinduced generation of superoxide [71, 89, 90]. Initially, melanosome degradation is accompanied by reduction in their capacity to bind iron ions, and to offer protection against iron ion-mediated decomposition of hydrogen peroxide and lipid peroxidation (Figure 7.11). Further degradation of melanosomes leads to a complete loss of inhibitory effect on oxidation catalyzed by metal ions. Instead, degraded melanosomes become pro-oxidant and mediate metal ion-induced oxidation. It remains to be investigated what changes in melanin structure are responsible for the switch from an anti- to pro-oxidant. It is tempting to suggest that RPE melanosomes may undergo degradation in vivo due to life-long exposure to light and oxidants and, as a result, lose their ability to protect the cells from oxidation catalyzed by metal ions or even become prooxidant and mediate metal ion-induced oxidation. This could have profound consequences for the retina where the impairment in iron transport and metabolism, and accumulation of excessive amounts of iron, is associated with several retinal degenerations [91, 92]. In the aging RPE, oxidative changes in melanin may increase due to formation of complex granules with lipofuscin, which can generate ROS upon excitation with

213

214

7 Properties and Functions of Ocular Melanins and Melanosomes a)

Effect of photodegradation of human RPE melanosomes on oxygen uptake induced by Fe-ADP/ascorbate (0.05 mM/0.2 mM) in the absence (a) and the presence (b) of 10 mM linoleate (LA). To induce degradation, melanosomes were

Figure 7.11

b)

exposed to 0.95 or 5.54 kJ/cm2 of blue light delivered from a compact arc high-pressure mercury lamp equipped with broadband ﬁlter transmitting 390- to 490-nm light. (Modiﬁed from [80].)

blue light [3, 86]. Moreover, the complex granules of melanolipofuscin are 2.8- and 2.1-fold more susceptible to oxidation induced by iron ions than melanosomes or lipofuscin, respectively, isolated from the same donors (Figure 7.12). 7.7.3 Cytotoxic Properties of Aged RPE Melanin Granules and Their Potential Consequences for Retinal Aging and AMD

Deleterious effects of aged human melanosomes and melanolipofuscin have been demonstrated in the cultured RPE cell line ARPE-19 exposed to blue/green light (Figure 7.13) [49, 110]. The cells were enriched with bovine and human RPE melanosomes and human melanolipofuscin by feeding. Due to differences in the experimental conditions used in these two studies, the comparison of the effects of melanosomes and melanolipofuscin on susceptibility to light-induced toxicity can only be made indirectly. Exposure to light of cells enriched with bovine melanosomes led to a small decrease of 11% of mitochondrial activity. Human melanosomes isolated from donors aged 60–90 years induced death of about 44% of cells, whereas exposure of lipofuscin-laden cells under the same conditions led to death of about 70% cells. Exposure of melanolipofuscin-laden cells to blue/ green light induced death of about 58% of cells, whereas exposure of lipofuscinladen cells under the same conditions led to death of 80% cells. These data demonstrate that aged melanosomes and melanolipofuscin exhibit substantial phototoxicity. Considering that in elderly people melanolipofuscin is the predomi-

7.7 Pro-Oxidant Properties of Ocular Melanosomes

Figure 7.12 Comparison of the rates of oxygen uptake induced by Fe-ADP/ascorbate (0.05 mM/0.2 mM) in suspensions of bovine (BMs) and human (HMs) RPE melanosomes, melanolipofuscin (HMLf), and lipofuscin (HLf). (Modiﬁed from [80].)

Figure 7.13 Mitochondrial activity of ARPE-19 cells without (Control) and with internalized human RPE melanosomes (OHMs) or bovine melanosomes (BMs) as a

function of time of exposure to blue/green light (390–550 nm; 2.8 mW/cm2). Human RPE melanosomes were isolated from 60- to 90-year-old donors. (Modiﬁed from [110].)

nant pigment granule in the RPE, its properties with regard to interaction with light and redox-active metal ions require further investigation. The properties of RPE melanosomes and their changes that occur with age suggest that RPE melanosomes may play a role in retinal aging and diseases, such as AMD. Aging of the RPE is accompanied by partial loss of melanin, and accumulation of melanolipofuscin and iron. It can be suggested that with aging the antioxidant properties of RPE melanosomes become gradually outbalanced by their pro-oxidant properties. Due to these pro-oxidant properties RPE melanosomes may play a causal role in the impairment of iron homeostasis and the increase of oxidative stress in the RPE. There is a growing body of evidence

215

216

7 Properties and Functions of Ocular Melanins and Melanosomes

pointing to the RPE as the primary site of damage leading to AMD. The characteristic features of AMD include increased accumulation of iron in the RPE and several markers of oxidative damage. Several products resulting from oxidative damage to constituents that accumulate in the retina can trigger a vicious cycle involving proinﬂammatory and proangiogenic pathways, which results in more oxidative stress in the retina and further damage. Thus, understanding the agerelated changes in structure and properties of RPE melanosomes may help to understand the initial pathways leading to RPE dysfunction followed by development of AMD.

7.8 Other Properties of Ocular Melanosomes and Their Implications

Apart from the anti- and pro-oxidant properties discussed above, ocular melanosomes exhibit several other characteristics that are important for the eye, starting from early embryogenesis. Proper maturation of melanosomes is required for normal development of the visual system [6]. Due to the congenital reduction or absence of melanin synthesis in the eye, not only does hypopigmentation of ocular structures occur, but the retina remains underdeveloped with an absent fovea and there are misguided neurons in the optic chiasma. The defects associated with ocular albinism manifest clinically as lack of acute vision, nystagmus, and strabismus. In addition to antimalarial drugs mentioned previously, there are a number of other xenobiotics with high afﬁnity for binding to melanin, including drugs for rheumatoid arthritis, local anesthetics, aminoglycoside antibiotics, and phenothiazine-based drugs such as chlorpromazine used as antipsychotics [84, 111]. Moreover, some environmental pollutants including herbicides, dyes, and alkaloids also bind to melanin. Some types of bonds between melanin and xenobiotics are ionic and easily reversible, but irreversible binding was observed with chloroquine and chlorpromazine where autoradiographic study of [14C]chloroquine and [35S]chlorpromazine revealed that radioactivity in pigmented structures persisted for up to 90 days and 1 year for [14C]chloroquine and [35S]chlorpromazine, respectively. On one hand, binding of xenobiotics to melanin may be protective when the noxious chemicals are rapidly bound to melanin and slowly released at small nontoxic concentrations [112]. It may be particularly important for the retina where the RPE provides the blood–retina barrier so potentially toxic substances may become bound to RPE melanin and therefore disabled from reaching photoreceptors. Adverse effects of binding of drugs to melanin include accumulation of drugs in pigmented tissues, leading eventually to degenerative changes and toxicity such as those observed in the RPE. Binding of drugs to melanin can also affect ocular drug delivery [113]. Ocular melanosomes accumulate considerable amounts of metals such as calcium and zinc [84, 114, 115]. It has been suggested that binding of calcium to

7.9 Conclusions

melanin may play a role in calcium regulation and buffering, which seem to be particularly important for cell adhesion and for retinal photoreceptors with tightly regulated calcium levels. Interestingly, the concentration of calcium in ocular pigmented tissues is very high, exceeding even that of the bone where calcium is deposited in a mineral form [116–118]. X-ray microanalyses of elemental compositions of choroidal and RPE melanosomes from an 8-month-old rat, 2-year-old Cynomolgus monkey and 68-year-old human show that calcium accounts for 0.04 to 0.78% mol fraction. X-ray microanalyses of ﬁxed sections of ocular tissues of various species have demonstrated that melanosomes of the ciliary body, iris, RPE, and choroid contain up to 2–10 times more calcium than adjacent nonpigmented cellular organelles [40, 118]. Choroidal melanosomes of the rat, rabbit, pig, and monkey contain up to 7-fold greater concentrations of calcium than RPE melanosomes [40]. RPE melanosomes contain cathepsin D, which is the main hydrolase responsible for the proteolysis of opsin from phagocytosed photoreceptor outer segments [48]. The presence of cathepsin D at the surface of RPE melanosomes and observations of fusion between RPE melanosomes and phagosomes suggest that melanosomes may be involved in the degradation of photoreceptor outer segments. Support for this hypothesis comes also from experiments on cultured RPE cells fed daily for up to 4 weeks with photoreceptor outer segments [119]. Nonpigmented RPE cells isolated from the albino rabbit and nonpigmented tapetal part of bovine fundus showed greater accumulation of nondigested photoreceptor outer segments than pigmented rabbit and bovine RPE cells. Absorption of light by the RPE and choroidal melanosomes and their IR ﬂuorescence have been employed in the development of diagnostic tools for imaging of the retina–choroid complex [120]. Absorption of light together with an efﬁcient thermal deactivation pathway in melanosomes have also been employed in therapeutic photocoagulation of ocular tissues [121].

7.9 Conclusions

Undoubtedly, ocular melanosomes play multiple roles in the eye. The most obvious and well documented is their role as a broadband optical ﬁlter that, in the case of the iris, can play a protective role by limiting light intensity reaching the retina, crystalline lens, as well as the nuclei of iridial melanocytes. A potential photoprotective role of RPE and choroidal melanosomes still awaits experimental evidence. Aging of the RPE melanosomes leads to an increase in their pro-oxidant properties which may counterbalance or even exceed their antioxidant properties. This may increase the oxidative stress in the aged RPE and contribute to agerelated retinal dysfunction and development of AMD. It remains to be determined at the molecular level what structural changes of the RPE melanin/melanosomes are responsible for the observed changes in their morphology and physicochemical properties.

217

218

7 Properties and Functions of Ocular Melanins and Melanosomes

References 1 Imesch, P.D., Bindley, C.D., Khademian,

11 Jimbow, K., Hara, H., Vinayagamoorthy,

Z., Ladd, B., Gangnon, R., Albert, D.M., and Wallow, I.H. (1996) Melanocytes and iris color. Electron microscopic ﬁndings. Arch. Ophthalmol., 114, 443–447. Hu, D.N., Simon, J.D., and Sarna, T. (2008) Role of ocular melanin in ophthalmic physiology and pathology. Photochem. Photobiol., 84, 639–644. Rozanowska, M., Rozanowski, B., and Boulton, M. (2009) Photobiology of the retina: light-induced damage to the retina, in Photobiological Sciences Online (ed. K.C. Smith), American Society for Photobiology, Lawrence, KS, http:// www.photobiology.info/ Rozanowska.html. Sarna, T. (1992) Properties and function of the ocular melanin – a photobiophysical view. J. Photochem. Photobiol. B, 12, 215–258. Schraermeyer, U. and Heimann, K. (1999) Current understanding on the role of retinal pigment epithelium and its pigmentation. Pigment Cell Res., 12, 219–236. Boulton, M. (1998) Melanin and the retinal pigment epithelium, in The Retinal Pigment Epithelium: Function and Disease (eds M.F. Marmor and T.J. Wolfensberger), Oxford University Press, New York, pp. 68–85. Giebel, L.B. and Spritz, R.A. (1992) The molecular basis of type I (tyrosinasedeﬁcient) human oculocutaneous albinism. Pigment Cell Res., (Suppl. 2), 101–106. Jackson, I.J., Chambers, D.M., Tsukamoto, K., Copeland, N.G., Gilbert, D.J., Jenkins, N.A., and Hearing, V. (1992) A second tyrosinase-related protein, TRP-2, maps to and is mutated at the mouse slaty locus. EMBO J., 11, 527–535. Shibahara, S. (1993) Functional analysis of the tyrosinase gene and brown-locus protein gene promoters. J. Invest. Dermatol., 100, 146S–149S. Spritz, R.A. (1993) Molecular genetics of oculocutaneous albinism. Semin. Dermatol., 12, 167–172.

T., Luo, D., Dakour, J., Yamada, K., Dixon, W., and Chen, H. (1994) Molecular control of melanogenesis in malignant melanoma: functional assessment of tyrosinase and lamp gene families by UV exposure and gene co-transfection, and cloning of a cDNA encoding calnexin, a possible melanogenesis “chaperone”. J. Dermatol., 21, 894–906. Simon, J.D., Peles, D., Wakamatsu, K., and Ito, S. (2009) Current challenges in understanding melanogenesis: bridging chemistry, biological control, morphology, and function. Pigment Cell Melanoma Res., 22, 563–579. Ito, S. and Wakamatsu, K. (2008) Chemistry of mixed melanogenesis – pivotal roles of dopaquinone. Photochem. Photobiol., 84, 582–592. Fuller, B.B., Spaulding, D.T., and Smith, D.R. (2001) Regulation of the catalytic activity of preexisting tyrosinase in black and Caucasian human melanocyte cell cultures. Exp. Cell Res., 262, 197–208. Smith, D.R., Spaulding, D.T., Glenn, H.M., and Fuller, B.B. (2004) The relationship between Na+/H+ exchanger expression and tyrosinase activity in human melanocytes. Exp Cell Res, 298, 521–534. Cheli, Y., Luciani, F., Khaled, M., Beuret, L., Bille, K., Gounon, P., Ortonne, J.P., Bertolotto, C., and Ballotti, R. (2009) αMSH and cyclic AMP elevating agents control melanosome pH through a protein kinase A-independent mechanism. J. Biol. Chem., 284, 18699–18706. Dryja, T.P., O’Neil-Dryja, M., Pawelek, J.M., and Albert, D.M. (1978) Demonstration of tyrosinase in the adult bovine uveal tract and retinal pigment epithelium. Invest. Ophthalmol. Vis. Sci., 17, 511–514. Nakazawa, M., Tsuchiya, M., Hayasaka, S., and Mizuno, K. (1985) Tyrosinase activity in the uveal tissue of the adult bovine eye. Exp. Eye Res., 41, 249–258.

2

3

4

5

6

7

8

9

10

12

13

14

15

16

17

18

References 19 Hayasaka, S., Nakazawa, M., Ishiguro,

20

21

22

23

24

25

26

27

28

29

30

S., and Mizuno, K. (1986) Presence of tyrosinase activity in human ciliary body. Jpn. J. Ophthalmol., 30, 32–35. Feeney-Burns, L., Hilderbrand, E.S., and Eldridge, S. (1984) Aging human RPE: morphometric analysis of macular, equatorial, and peripheral cells. Invest. Ophthalmol. Vis. Sci., 25, 195–200. Weiter, J.J., Delori, F.C., Wing, G.L., and Fitch, K.A. (1986) Retinal pigment epithelial lipofuscin and melanin and choroidal melanin in human eyes. Invest. Ophthalmol. Vis. Sci., 27, 145–152. Schmidt, S.Y. and Peisch, R.D. (1986) Melanin concentration in normal human retinal pigment epithelium. Regional variation and age-related reduction. Invest. Ophthalmol. Vis. Sci., 27, 1063–1067. Sarna, T., Burke, J.M., Korytowski, W., Rozanowska, M., Skumatz, C.M., Zareba, A., and Zareba, M. (2003) Loss of melanin from human RPE with aging: possible role of melanin photooxidation. Exp. Eye Res., 76, 89–98. Biesemeier, A., Kreppel, F., Kochanek, S., and Schraermeyer, U. (2010) The classical pathway of melanogenesis is not essential for melanin synthesis in the adult retinal pigment epithelium. Cell Tissue Res., 339, 551–560. Hayasaka, S. (1989) Aging changes in lipofuscin, lysosomes and melanin in the macular area of human retina and choroid. Jpn. J. Ophthalmol., 33, 36–42. Feeney-Burns, L., Burns, R.P., and Gao, C.L. (1990) Age-related macular changes in humans over 90 years old. Am. J. Ophthalmol., 109, 265–278. Hunold, W. and Malessa, P. (1974) Spectrophotometric determination of the melanin pigmentation of the human ocular fundus in vivo. Ophthalmic Res., 6, 355–362. Van Norren, D. and Tiemeijer, L.F. (1986) Spectral reﬂectance of the human eye. Vision Res., 26, 313–320. Delori, F.C. and Pﬂibsen, K.P. (1989) Spectral reﬂectance of the human ocular fundus. Appl. Opt., 28, 1061–1077. van de Kraats, J., Berendschot, T.T., and van Norren, D. (1996) The pathways of

31

32

33

34

35

36

37

38

39

40

light measured in fundus reﬂectometry. Vision Res., 36, 2229–2247. Prota, G., Hu, D.N., Vincensi, M.R., McCormick, S.A., and Napolitano, A. (1998) Characterization of melanins in human irides and cultured uveal melanocytes from eyes of different colors. Exp. Eye Res., 67, 293–299. Wakamatsu, K., Hu, D.N., McCormick, S.A., and Ito, S. (2008) Characterization of melanin in human iridal and choroidal melanocytes from eyes with various colored irides. Pigment Cell Melanoma Res., 21, 97–105. Wielgus, A.R. and Sarna, T. (2005) Melanin in human irides of different color and age of donors. Pigment Cell Res., 18, 454–464. Cracknell, K.P. and Grierson, I. (2009) Prostaglandin analogues in the anterior eye: their pressure lowering action and side effects. Exp. Eye Res., 88, 786–791. Prota, G., Vincensi, M.R., Napolitano, A., Selen, G., and Stjernschantz, J. (2000) Latanoprost stimulates eumelanogenesis in iridial melanocytes of cynomolgus monkeys. Pigment Cell Res., 13, 147–150. Hu, F. and Mah, K. (1983) Changes in melanosomes with age in iridial stromal melanocytes of rhesus macaques. Mech. Ageing Dev., 23, 95–102. Hu, D.N., McCormick, S.A., Ritch, R., and Pelton-Henrion, K. (1993) Studies of human uveal melanocytes in vitro: isolation, puriﬁcation and cultivation of human uveal melanocytes. Invest. Ophthalmol. Vis. Sci., 34, 2210–2219. Hu, D.N., Ritch, R., McCormick, S.A., and Pelton-Henrion, K. (1992) Isolation and cultivation of human iris pigment epithelium. Invest. Ophthalmol. Vis. Sci., 33, 2443–2453. Liu, Y., Hong, L., Wakamatsu, K., Ito, S., Adhyaru, B.B., Cheng, C.Y., Bowers, C.R., and Simon, J.D. (2005) Comparisons of the structural and chemical properties of melanosomes isolated from retinal pigment epithelium, iris and choroid of newborn and mature bovine eyes. Photochem. Photobiol., 81, 510–516. Eibl, O., Schultheiss, S., BlitgenHeinecke, P., and Schraermeyer, U.

219

220

7 Properties and Functions of Ocular Melanins and Melanosomes

41

42

43

44

45

46

47

48

(2006) Quantitative chemical analysis of ocular melanosomes in the TEM. Micron, 37, 262–276. Hu, F. and Mah, K. (1979) Choroidal melanocytes – a model for studying the aging process in nonreplicative differentiated cells. Mech. Ageing Dev., 11, 227–235. Feeney, L. (1978) Lipofuscin and melanin of human retinal pigment epithelium. Fluorescence, enzyme cytochemical, and ultrastructural studies. Invest. Ophthalmol. Vis. Sci., 17, 583–600. Ward, W.C. and Simon, J.D. (2007) The differing embryonic origins of retinal and uveal (iris/ciliary body and choroid) melanosomes are mirrored by their phospholipid composition. Pigment Cell Res., 20, 61–69. Peles, D.N., Hong, L., Hu, D.N., Ito, S., Nemanich, R.J., and Simon, J.D. (2009) Human iridal stroma melanosomes of varying pheomelanin contents possess a common eumelanic outer surface. J. Phys. Chem. B, 113, 11346–11351. Brumbaugh, J.A. (1968) Ultrastructural differences between forming eumelanin and pheomelanin as revealed by the pink-eye mutation in the fowl. Dev. Biol., 18, 375–390. Samokhvalov, A., Hong, L., Liu, Y., Garguilo, J., Nemanich, R.J., Edwards, G.S., and Simon, J.D. (2005) Oxidation potentials of human eumelanosomes and pheomelanosomes. Photochem. Photobiol., 81, 145–148. Ye, T., Hong, L., Garguilo, J., Pawlak, A., Edwards, G.S., Nemanich, R.J., Sarna, T., and Simon, J.D. (2006) Photoionization thresholds of melanins obtained from free electron laserphotoelectron emission microscopy, femtosecond transient absorption spectroscopy and electron paramagnetic resonance measurements of oxygen photoconsumption. Photochem. Photobiol., 82, 733–737. Azarian, S.M., McLeod, I., Lillo, C., Gibbs, D., Yates, J.R., and Williams, D.S. (2006) Proteomic analysis of mature melanosomes from the retinal pigmented epithelium. J. Proteome Res., 5, 521–529.

49 Warburton, S., Davis, W.E., Southwick,

50

51

52

53

54

55

56

57

58

K., Xin, H., Woolley, A.T., Burton, G.F., and Thulin, C.D. (2007) Proteomic and phototoxic characterization of melanolipofuscin: correlation to disease and model for its origin. Mol. Vis., 13, 318–329. Ng, K.P., Gugiu, B., Renganathan, K., Davies, M.W., Gu, X., Crabb, J.S., Kim, S.R., Rozanowska, M.B., Bonilha, V.L., Rayborn, M.E., Salomon, R.G., Sparrow, J.R., Boulton, M.E., Hollyﬁeld, J.G., and Crabb, J.W. (2008) Retinal pigment epithelium lipofuscin proteomics. Mol. Cell Proteomics, 7, 1397–1405. Peles, D.N. and Simon, J.D. (2010) Direct measurement of the ultraviolet absorption coefﬁcient of single retinal melanosomes. Photochem. Photobiol., 86, 279–281. Meredith, P. and Sarna, T. (2006) The physical and chemical properties of eumelanin. Pigment Cell Res., 19, 572–594. Sarna, T. and Rozanowska, M. (1994) Phototoxicity to the eye, in Photobiology in Medicine (eds G. Jori, R.H. Pottier, M.A.J. Rodgers, and T.G. Truscott), Plenum Press, New York, pp. 125–142. Hashemi, H., Yazdani, K., Khabazkhoob, M., Mehravaran, S., Mohammad, K., and Fotouhi, A. (2009) Distribution of photopic pupil diameter in the Tehran Eye Study. Curr. Eye Res., 34, 378–385. Watts, G.K. (1971) Retinal hazards during laser irradiation of the iris. Br. J. Ophthalmol., 55, 60–67. Hashemi, H., KhabazKhoob, M., Yazdani, K., Mehravaran, S., Mohammad, K., and Fotouhi, A. (2010) White-to-white corneal diameter in the Tehran Eye Study. Cornea, 29, 9–12. IJspeert, J.K., de Waard, P.W., van den Berg, T.J., and de Jong, P.T. (1990) The intraocular straylight function in 129 healthy volunteers; dependence on angle, age and pigmentation. Vision Res., 30, 699–707. van den Berg, T.J., IJspeert, J.K., and de Waard, P.W. (1991) Dependence of intraocular straylight on pigmentation and light transmission through the ocular wall. Vision Res., 31, 1361–1367.

References 59 Hashemi, H., KhabazKhoob, M., Yekta,

60

61

62

63

64

65

66

67

68

A., Mohammad, K., and Fotouhi, A. (2010) Distribution of iris colors and its association with ocular disorder in the Tehran Eye Study. Iranian J. Ophthalmol., 22, 7–14. Hetherington, A.M. and Johnson, B.E. (1984) Photohemolysis. Photodermatology, 1, 255–260. Misra, R.B., Ray, R.S., and Hans, R.K. (2005) Effect of UVB radiation on human erythrocytes in vitro. Toxicol. In Vitro, 19, 433–438. Wu, X., Pan, L., Wang, Z., Liu, X., Zhao, D., Zhang, X., Rupp, R.A., and Xu, J. (2010) Ultraviolet irradiation induces autoﬂuorescence enhancement via production of reactive oxygen species and photodecomposition in erythrocytes. Biochem. Biophys. Res. Commun., 396, 999–1005. Ankri, R., Friedman, H., Savion, N., Kotev-Emeth, S., Breitbart, H., and Lubart, R. (2010) Visible light induces nitric oxide (NO) formation in sperm and endothelial cells. Lasers Surg. Med., 42, 348–352. Lavi, R., Shainberg, A., Shneyvays, V., Hochauser, E., Isaac, A., Zinman, T., Friedmann, H., and Lubart, R. (2010) Detailed analysis of reactive oxygen species induced by visible light in various cell types. Lasers Surg. Med., 42, 473–480. Hu, D.N. (2005) Photobiology of ocular melanocytes and melanoma. Photochem. Photobiol., 81, 506–509. Sarna, T. and Swartz, H.M. (2006) The physical properties of melanins, in The Pigmentary System: Physiology and Pathophysiology (eds J.J. Nordlund, R.E. Boissy, V.J. Hearing, R.A. King, and J.-P. Ortonne), Blackwell, Malden, MA, pp. 305–335. Rozanowska, M., Sarna, T., Land, E.J., and Truscott, T.G. (1999) Free radical scavenging properties of melanin interaction of eu- and pheo-melanin models with reducing and oxidising radicals. Free Radic. Biol. Med., 26, 518–525. Serpentini, C.L., Gauchet, C., de Montauzon, D., Comtat, M., Ginestar, J., and Paillous, N. (2000) First

69

70

71

72

73

74

75

76

77

electrochemical investigation of the redox properties of DOPA-melanins by means of a carbon paste electrode. Electrochim. Acta, 45, 1663–1668. Edge, R., Land, E.J., Rozanowska, M., Sarna, T., and Truscott, T.G. (2000) Carotenoid radical–melanin interactions. J. Phys. Chem. B, 104, 7193–7196. Halliwell, B. and Gutteridge, J.M.C. (1998) Free Radicals in Biology and Medicine, Oxford University Press, New York. Rozanowska, M., Korytowski, W., Rozanowski, B., Skumatz, C., Boulton, M.E., Burke, J.M., and Sarna, T. (2002) Photoreactivity of aged human RPE melanosomes: a comparison with lipofuscin. Invest. Ophthalmol. Vis. Sci., 43, 2088–2096. Miceli, M.V., Liles, M.R., and Newsome, D.A. (1994) Evaluation of oxidative processes in human pigment epithelial cells associated with retinal outer segment phagocytosis. Exp. Cell Res., 214, 242–249. Augusto, O., Bonini, M.G., Amanso, A.M., Linares, E., Santos, C.C., and De Menezes, S.L. (2002) Nitrogen dioxide and carbonate radical anion: two emerging radicals in biology. Free Radic. Biol. Med., 32, 841–859. Stepien, K., Wilczok, A., Zajdel, A., Dzierzega-Lecznar, A., and Wilczok, T. (2000) Peroxynitrite mediated linoleic acid oxidation and tyrosine nitration in the presence of synthetic neuromelanins. Acta Biochim. Pol., 47, 931–940. Stepien, K., Zajdel, A., Wilczok, A., Wilczok, T., Grzelak, A., Mateja, A., Soszynski, M., and Bartosz, G. (2000) Dopamine-melanin protects against tyrosine nitration, tryptophan oxidation and Ca2+-ATPase inactivation induced by peroxynitrite. Biochim. Biophys. Acta, 1523, 189–195. Miyagi, M., Sakaguchi, H., Darrow, R.M., Yan, L., West, K.A., Aulak, K.S., Stuehr, D.J., Hollyﬁeld, J.G., Organisciak, D.T., and Crabb, J.W. (2002) Evidence that light modulates protein nitration in rat retina. Mol. Cell Proteomics, 1, 293–303. Rózanowska, M. and Rózanowski, B. (2010) Uptake and photoprotection in

221

222

7 Properties and Functions of Ocular Melanins and Melanosomes

78

79

80

81

82

83

84

85

86

cultured RPE cells, in Carotenoids: Physical, Chemical, and Biological Functions and Properties (ed. J.T. Landrum), CRC Press, Boca Raton, FL, pp. 309–364. Dunford, R., Land, E.J., Rozanowska, M., Sarna, T., and Truscott, T.G. (1995) Interaction of melanin with carbon- and oxygen-centered radicals from methanol and ethanol. Free Radic. Biol. Med., 19, 735–740. Ye, T., Simon, J.D., and Sarna, T. (2003) Ultrafast energy transfer from bound tetra(4-N,N,N,N-trimethylanilinium) porphyrin to synthetic dopa and cysteinyldopa melanins. Photochem. Photobiol., 77, 1–4. Rozanowski, B., Burke, J.M., Boulton, M.E., Sarna, T., and Rozanowska, M. (2008) Human RPE melanosomes protect from photosensitized and iron-mediated oxidation but become pro-oxidant in the presence of iron upon photodegradation. Invest. Ophthalmol. Vis. Sci., 49, 2838–2847. Sarna, T., Rozanowska, M., Zareba, M., Korytowski, W., and Boulton, M. (1998) Retinal melanin and lipofuscin: possible role in photoprotection and phototoxicity of the human RPE. 12th International Congress on Photobiology, Vienna Spikes, J.D. (1998) Photosensitizing properties of quinine and synthetic antimalarials. J. Photochem. Photobiol. B, 42, 1–11. Motten, A.G., Martinez, L.J., Holt, N., Sik, R.H., Reszka, K., Chignell, C.F., Tonnesen, H.H., and Roberts, J.E. (1999) Photophysical studies on antimalarial drugs. Photochem. Photobiol., 69, 282–287. Larsson, B.S. (1993) Interaction between chemicals and melanin. Pigment Cell Res., 6, 127–133. Dewi, N.A., Yuzawa, M., Tochigi, K., Kawamura, A., and Mori, R. (2008) Effects of photodynamic therapy on the choriocapillaris and retinal pigment epithelium in the irradiated area. Jpn. J. Ophthalmol., 52, 277–281. Rozanowska, M. and Rozanowski, B. (2008) Visual transduction and age-related changes in lipofuscin, in Visual Transduction and Non-Visual

87

88

89

90

91

92

93

94

95

Perception (eds J. Tombran-Tink and C.J. Barnstable), Humana Press, Totowa, NJ, pp. 405–446. Wang, A., Marino, A.R., Gasyna, Z., Gasyna, E., and Norris, J., Jr (2008) Photoprotection by porcine eumelanin against singlet oxygen production. Photochem. Photobiol., 84, 679–682. Hong, L. and Simon, J.D. (2007) Current understanding of the binding sites, capacity, afﬁnity, and biological signiﬁcance of metals in melanin. J. Phys. Chem. B, 111, 7938–7947. Zadlo, A., Rozanowska, M.B., Burke, J.M., and Sarna, T.J. (2007) Photobleaching of retinal pigment epithelium melanosomes reduces their ability to inhibit iron-induced peroxidation of lipids. Pigment Cell Res., 20, 52–60. Zareba, M., Szewczyk, G., Sarna, T., Hong, L., Simon, J.D., Henry, M.M., and Burke, J.M. (2006) Effects of photodegradation on the physical and antioxidant properties of melanosomes isolated from retinal pigment epithelium. Photochem. Photobiol., 82, 1024–1029. Wong, R.W., Richa, D.C., Hahn, P., Green, W.R., and Dunaief, J.L. (2007) Iron toxicity as a potential factor in AMD. Retina, 27, 997–1003. He, X., Hahn, P., Iacovelli, J., Wong, R., King, C., Bhisitkul, R., MassaroGiordano, M., and Dunaief, J. (2007) Iron homeostasis and toxicity in retinal degeneration. Prog. Retin. Eye Res., 26, 649–673. Hahn, P., Ying, G.S., Beard, J., and Dunaief, J.L. (2006) Iron levels in human retina: sex difference and increase with age. Neuroreport, 17, 1803–1806. Hahn, P., Milam, A.H., and Dunaief, J.L. (2003) Maculas affected by age-related macular degeneration contain increased chelatable iron in the retinal pigment epithelium and Bruch’s membrane. Arch. Ophthalmol., 121, 1099–1105. Liles, M.R., Newsome, D.A., and Oliver, P.D. (1991) Antioxidant enzymes in the aging human retinal pigment

References

96

97

98

99

100

101

102

103

104

105

epithelium. Arch. Ophthalmol., 109, 1285–1288. Boulton, M., Rozanowska, M., and Rozanowski, B. (2001) Retinal photodamage. J. Photochem. Photobiol. B, 64, 144–161. Seagle, B.L., Rezai, K.A., Kobori, Y., Gasyna, E.M., Rezaei, K.A., and Norris, J.R., Jr (2005) Melanin photoprotection in the human retinal pigment epithelium and its correlation with light-induced cell apoptosis. Proc. Natl. Acad. Sci. USA, 102, 8978–8983. Zareba, M., Raciti, M.W., Henry, M.M., Sarna, T., and Burke, J.M. (2006) Oxidative stress in ARPE-19 cultures: do melanosomes confer cytoprotection? Free Radic. Biol. Med., 40, 87–100. Rozanowska, M., Bober, A., Burke, J.M., and Sarna, T. (1997) The role of retinal pigment epithelium melanin in photoinduced oxidation of ascorbate. Photochem. Photobiol., 65, 472–479. Grzelak, A., Rychlik, B., and Bartosz, G. (2001) Light-dependent generation of reactive oxygen species in cell culture media. Free Radic. Biol. Med., 30, 1418–1425. Rozanowska, M., Jarvisevans, J., Korytowski, W., Boulton, M.E., Burke, J.M., and Sarna, T. (1995) Blue light-induced reactivity of retinal age pigment – in-vitro generation of oxygen-reactive species. J. Biol. Chem., 270, 18825–18830. Rózanowska, M. (1998) Badania fotoreaktywności in vitro komórek nabłonka upigmentowanego siatkówki. PhD thesis. Department of Biophysics, Institute of Molecular Biology, Jagiellonian University, Kraków, Poland. Dontsov, A.E., Glickman, R.D., and Ostrovsky, M.A. (1999) Retinal pigment epithelium pigment granules stimulate the photo-oxidation of unsaturated fatty acids. Free Radic. Biol. Med., 26, 1436–1446. Wielgus, A.R. and Sarna, T. (2008) Ascorbate enhances photogeneration of hydrogen peroxide mediated by the iris melanin. Photochem. Photobiol., 84, 683–691. Rozanowski, B., Burke, J., Sarna, T., and Rozanowska, M. (2008) The pro-oxidant

106

107

108

109

110

111

112

113

114

115

effects of interactions of ascorbate with photoexcited melanin fade away with aging of the retina. Photochem. Photobiol., 84, 658–670. Glickman, R.D. and Lam, K.W. (1992) Oxidation of ascorbic acid as an indicator of photooxidative stress in the eye. Photochem. Photobiol., 55, 191–196. Glickman, R.D., Sowell, R., and Lam, K.W. (1993) Kinetic properties of light-dependent ascorbic acid oxidation by melanin. Free Radic. Biol. Med., 15, 453–457. Sarna, T. and Swartz, H.M. (1998) The physical properties of melanins, in Pigmentary System and its Disorders (eds J.J. Nordlund, R.E. Boissy, V.J. Hearing, R.A. King, and J. Ortonne), Oxford University Press, New York, pp. 333–357. Kayatz, P., Thumann, G., Luther, T.T., Jordan, J.F., Bartz-Schmidt, K.U., Esser, P.J., and Schraermeyer, U. (2001) Oxidation causes melanin ﬂuorescence. Invest. Ophthalmol. Vis. Sci., 42, 241–246. Rozanowski, B., Cuenco, J., Davies, S., Shamsi, F.A., Zadlo, A., Dayhaw-Barker, P., Rozanowska, M., Sarna, T., and Boulton, M. (2008) The phototoxicity of aged human retinal melanosomes. Photochem. Photobiol., 84, 650–657. Buszman, E., Beberok, A., Rozanska, R., and Orzechowska, A. (2008) Interaction of chlorpromazine, ﬂuphenazine and triﬂuoperazine with ocular and synthetic melanin in vitro. Pharmazie, 63, 372–376. Zemel, E., Loewenstein, A., Lei, B., Lazar, M., and Perlman, I. (1995) Ocular pigmentation protects the rabbit retina from gentamicin-induced toxicity. Invest. Ophthalmol. Vis. Sci., 36, 1875–1884. Gaudana, R., Ananthula, H.K., Parenky, A., and Mitra, A.K. (2010) Ocular drug delivery. AAPS J., 12, 348–360. Kokkinou, D., Kasper, H.U., BartzSchmidt, K.U., and Schraermeyer, U. (2004) The pigmentation of human iris inﬂuences the uptake and storing of zinc. Pigment Cell Res., 17, 515–518. Kokkinou, D., Kasper, H.U., Schwarz, T., Bartz-Schmidt, K.U., and Schraermeyer, U. (2005) Zinc uptake

223

224

7 Properties and Functions of Ocular Melanins and Melanosomes

116

117

118

119

and storage: the role of fundus pigmentation. Graefes Arch. Clin. Exp. Ophthalmol., 243, 1050–1055. Drager, U.C. (1985) Calcium binding in pigmented and albino eyes. Proc. Natl. Acad. Sci. USA, 82, 6716–6720. Drager, U.C. and Balkema, G.W. (1987) Does melanin do more than protect from light? Neurosci. Res. Suppl., 6, S75–S86. Panessa, B.J. and Zadunaisky, J.A. (1981) Pigment granules: a calcium reservoir in the vertebrate eye. Exp. Eye Res., 32, 593–604. Sundelin, S.P., Nilsson, S.E., and Brunk, U.T. (2001) Lipofuscin formation in

cultured retinal pigment epithelial cells is related to their melanin content. Free Radic. Biol. Med., 30, 74–81. 120 Keilhauer, C.N. and Delori, F.C. (2006) Near-infrared autoﬂuorescence imaging of the fundus: visualization of ocular melanin. Invest. Ophthalmol. Vis. Sci., 47, 3556–3564. 121 Stefansson, E. (2001) The therapeutic effects of retinal laser treatment and vitrectomy. A theory based on oxygen and vascular physiology. Acta Ophthalmol. Scand., 79, 435–440.

225

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease Kay L. Double, Wakako Maruyama, Makoko Naoi, Manfred Gerlach, and Peter Riederer

8.1 What are Neuromelanins?

Neuromelanins are one of the two primary pigment molecules in the human brain, the other being the so-called “age pigment” lipofuscin. No formal criteria to deﬁne a neuromelanin have been agreed upon, but a “neuromelanin” is generally considered to consist of a dark-colored, highly insoluble intracellular granular pigment found in the brain. In the human brain, neuromelanin granules have an amorphic shape, ranging in size from 0.5 to 2.5 μm and, because of their characteristic dark brown color, can readily be seen both macroscopically and at the light microscope level without the need for histochemical staining or immunohistochemical visualization (Figures 8.1a and 8.5a). Unlike lipofuscin, which is readily ﬂuorescent, neuromelanin does not immediate ﬂuoresce on exposure to UV light, although the pigment may develop this characteristic upon extended UV exposure [1]. In the healthy human brain the pigment is abundant in the cytoplasm of neurons [2] (Figure 8.1a), although it may also extend into cellular processes. Neuromelanin does not form in glial cells, but may sometimes be found in these cell types in the postmortem human brain in neurological disorders where pigmented neurons degenerate, as a result of the phagocytosis of the pigment released from dying cells. Unlike other melanins, at the electron microscope level neuromelanin granules exhibit a unique structure consisting of three components of different electron density (Figure 8.1b): an electron-dense melanin polymer, a component of intermediate electron density, and an electron-lucent lipid component not found in peripheral melanins or other nonpigmented central nervous system cells [3]. Individual pigment granules are heterogeneous in size and also in the proportion of each of the three components that they contain (Figure 8.1b). Further, we have observed large bodies of the electron-dense component of neuromelanin within neurons of the substantia nigra, although the function of these bodies is unknown [3]. Although a double membrane has been reported to surround granules of a synthetic neuromelanin formed in an in vitro system [4], there is no clear evidence for a membrane surrounding native neuromelanin granules in vivo in the human Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

226

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease a)

b)

Figure 8.1 (a) Neuromelanin-pigmented neuron from the human substantia nigra. The pigment appears dark under the light microscope and ﬁlls much of the cytoplasm. N, nucleus. (b) Appearance of neuromelanin granules under the electron microscope. The

pigment consists of three components of differing electron density: ED, electron-dense component; EI, electron intermediate component; EL, electron-lucent component. (Reproduced from [3].)

8.3 Development and Metabolism of Neuromelanin

brain [3, 5], suggesting that, unlike most other melanins, neuromelanins may interact directly with other molecules present in the cytoplasm.

8.2 Phylogenetic Development of Neuromelanin

Although there is a lack of studies speciﬁcally addressing this question, the presence of neuromelanin in the brain appears to be phylogenetically related to brain development. Studies of the midbrains of common laboratory and agricultural animals demonstrate an absence of the pigment in the substantia nigrae of lower species, such as mice, rats, and sheep. The pigment begins to ﬁrst appear in the brains of higher primate species, with small amounts of the pigment reported in the tyrosine hydroxylase (TH+)-positive neurons of the substantia nigra of primates, such as the baboon (Papio papio, Figure 8.2), although there have been no comprehensive comparative studies on the melanization of primate brains. The greatest quantify of the pigment, however, is clearly found in the human brain where the pigment ﬁlls 50% of the cytoplasm of TH+ neurons of the substantia nigra [2] (Figures 8.1 and 8.2), giving this brain region the characteristic dark appearance for which it is named (Figure 8.5a). Neuromelanin is also abundant in the neurons of the locus coeruleus, and is further found in the ventrolateral reticular formation and the nucleus of the solitary tract of the medulla oblongata (Figure 8.3). Neuromelanins appear to form from two of the three catecholamines – dopamine and norepinephrine, but not adrenalin [6, 7]. More recently, neuromelanins have been reported from other regions of the human brain such as the putamen, the premotor cortex, and the cerebellum [8]; however, as these brain regions do not exhibit dark pigments when viewed macro- or microscopically the nature of these putative neuromelanins is unknown.

8.3 Development and Metabolism of Neuromelanin

While the adult human midbrain contains the largest amounts of neuromelanin it is interesting that the development and subsequent appearance of the pigment is age-related. In fact, a dark neuromelanin is not present at all in the prenatal or infant human substantia nigra. Within the human substantia nigra, neuromelanin develops in three distinct stages [9]. The pigment ﬁrst appears in these neurons at approximately 3 years of age as small, pale granules, and the size and number of the granules increases until age 20, when the neuronal cytoplasm contains the adult complement of neuromelanin granules [9] (Figure 8.4). After the age of 20 the proportion of the neuron occupied by neuromelanin remains unaltered [9], but the amount of pigment, quantiﬁed biochemically, increases with increasing age [10], suggesting pigment granules are more compact in the aging brain,

227

228

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease a)

b)

c)

8.3 Development and Metabolism of Neuromelanin Cresyl violet-stained sections from the human (a), baboon (b), and rat (c) substantia nigrae. Human dopamine neurons contain the greatest quantify of neuromela-

Figure 8.2

nin pigment, while those of the baboon contain a relatively small amount of pigment and these neurons are unpigmented in the rat. (Reproduced from [18].)

a)

b)

c) Within the human brain, three main regions contain neuromelaninproducing cells: the substantia nigra of the midbrain (a), the locus coeruleus within the pons (b), and the ventrolateral reticular formation and the nucleus of the solitary

Figure 8.3

tract in the medulla oblongata (c). Of these regions, only the substantia nigra (a) and the locus coeruleus (b) contain a large cluster of pigmented neurons that can be seen macroscopically as a darkened area. (Reproduced from [18].)

possibly accounting for the observed darkening of the pigment with age [9]. Interestingly, the second common pigment in the human brain, lipofuscin, is sometimes referred to as the “age pigment,” yet the characteristic changes observed for neuromelanin with aging suggests this term could equally be applied to neuromelanin [3]. While the visual features of neuromelanin development in the human substantia nigra have now been described in detail, the pathways regulating the biosynthesis of the pigment remain largely unknown. The majority of melanins in the body are regulated via an enzymatic process, yet an enzyme that regulates neuromelanin synthesis is yet to be identiﬁed. The enzyme tyrosinase that controls melaninogenesis from the substrate l-tyrosine in other human tissues is absent in the substantia nigra and, despite a number of studies investigating this question, no other enzymatic regulatory pathway has been identiﬁed to date [9, 11]. The lack of a known regulatory system has resulted in a generalized belief that neuromelanin forms from an uncontrolled process of autoxidation of the

229

230

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease a)

3 years

d)

31 years

b)

8 years

e)

74 years

c)

19 years

f)

94 years

100µm Figure 8.4 Intermediate and high (inset) magniﬁcation photomicrographs of representative unstained neuromelaninpigmented neurons from the ventral region of the substantia nigra of humans at various ages. Scale bars in (f) are equivalent for all micrographs. The ﬁrst appearance of neuromelanin pigment is at 3 years of age

50µm

(a). The optical density of the neuromelanin pigment increases with age (a–f) while the average cellular volume occupied by neuromelanin pigment increases up to 20 years of age (a–c). After 20 the average cellular volume occupied by neuromelanin does not increase signiﬁcantly (d–f). (Reproduced from [9].)

8.4 Structure of Neuromelanin

catecholamines on which they are based. If this were true, the pigment would be expected to be present in approximately equal quantities in all catecholaminergic neurons of a given type. This is not the case, however, as, for example, 95% of the TH+ neurons of the substantia nigra are pigmented, while in the neighboring ventral tegmental area only 50% of TH+ neurons are pigmented [2]. Further, individuals medically treated with large quantities of l-3,4-dihydroxyphenylalanine (l-dopa), the precursor to dopamine, would be expected to exhibit increased levels of pigment within dopaminergic neurons as a result of increased autoxidation, but this is not the case [2]. The idea that neuromelanogenesis is nevertheless a controlled process is supported by the observation that nonpigmented fetal dopaminergic cells implanted into the striatal region of the brain as a treatment for Parkinson’s disease exhibit adult levels of pigmentation in just 3 years [12], suggesting that some yet to be identiﬁed factor or factors present in the adult brain support this process. Similarly, little is known about possible catabolism pathways for neuromelanin. Unlike pigmented cells in the periphery that may have a high rate of division and replacement, pigmented neurons in the brain do not divide. There is no known catabolism pathway for neuromelanin and, given the highly insoluble nature of the pigment, it is generally assumed that catabolism of the pigment, once formed, does not occur. Peripheral melanins are suggested to be catabolized via oxidative degradation and, interestingly, neuromelanin can be degraded under highly oxidative conditions in vitro. Recent biophysical data obtained from the pigment ex vivo demonstrates age-associated changes in highly oxidized forms of sulfur-based compounds in the pigment, suggesting that oxidative degradation may modify the mature pigment in the healthy human brain [13]. In disorders characterized by the death of the pigmented neurons of the substantia nigra, such as Parkinson’s disease and related disorders and toxin-induced substantia nigra cell death, the pigment is released by the dying neurons and can be seen within glial cells, which are assumed to remove and degrade the pigment from the brain [14, 15]. Changes in the appearance of neuromelanin in the human substantia nigra have been reported with aging, but carefully designed morphological studies suggest that the number of pigmented dopaminergic neurons in this brain region does not decrease with increasing age [16]. Nevertheless neuromelanin pigment is reported to be more commonly observed inside glial cells in the aged, compared with the young, brain [17].

8.4 Structure of Neuromelanin

As described above, neuromelanins form in some, but not all, catecholaminesynthesizing neurons and these neurotransmitters are considered to be primary substrates for the synthesis of the pigment. Thus, the neuromelanin of the substantia nigra is considered to be a dopamine-based melanin, while those of the locus coeruleus, reticular formation, and medulla oblongata are considered

231

232

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease

norepinephrine-based melanins. To date, the chemical structures of the norepinephrine-based neuromelanins are unknown, primarily because of the small amount of these melanins present in the human brain. The dopamine-based melanin of the human substantia nigra is relatively more abundant, but is very limited in availability because it occurs almost exclusively in the human brain. Nevertheless, studies on both synthetic melanin pigments and on native human neuromelanin have illuminated some of the structural features of this pigment. Neuromelanin is considered to be a copolymer of black insoluble eumelanin and the brown alkali-soluble pheomelanin, and shares structural features of both of these melanin types [18], with a polymer backbone based primarily upon l-dopa with reduced dopamine and dopamine metabolites also incorporated into the polymer [19], as well as a variety of reduced and oxidized sulfur species [13]. Interestingly, in contrast to the rat brain, which contains only one isoform of TH, the rate-limiting enzyme for dopamine production, the human brain involves four differentially regulated isoforms of this enzyme [20, 21]. This suggests that cellular levels of dopamine, and its metabolites, may be more variable in the human midbrain. Unlike other melanins, neuromelanin contains a signiﬁcant amount of associated lipids which make up 30% of the granule mass [22] (Figure 8.1b). The primary lipid associated with the melanin is the little-known isoprenoid dolichol with small amounts of other hydrophobic compounds such as cholesterol, ubiquinone, and ω-tocopherol also present [22, 23]. Amino acid analyses have demonstrated that a proteinaceous component comprises 5–15% of the pigment [19, 24]. In addition, proteomic analysis identiﬁed 72 proteins, some of which are associated with lysosomes and lysosome-related organelles [5, 25]. The apparent relative complexity of the structure of the pigment argues against the idea that the pigment is produced purely via a simple autoxidation pathway.

8.5 Biological Role of Neuromelanin in the Human Brain

For many years neuromelanin was considered to be an inert molecule with little or no physiological relevance, but more recent thinking suggests this view may not be correct. In tissues outside the central nervous system melanins are considered to have protective functions, the most obvious of which are the photoprotective functions of varying levels of melanin in the human skin, eye, and hair [18]. Neuromelanin may represent a cellular mechanism that evolved within catecholaminergic neurons to protect these cells from the potentially toxic metabolic products of these neurotransmitters. The enzymatic and autoxidation of dopamine, for example, produces highly oxidative quinone and semiquinone species that are suggested to be incorporated into the melanin polymer, effectively resulting in their inactivation [26–28]. Peripheral melanins have also been posited to be protective by virtue of their radical scavenging [29] and metal-binding properties [18, 30], and the free radical scavenging abilities of neuromelanins may represent an addi-

8.6 Is Neuromelanin Involved in Neurological Disease?

tional capability by which these molecules might beneﬁt the cell. We have demonstrated reduced cell death in the presence of neuromelanin in primary rat mesencephalic cultures challenged with an oxidative stimulus [31], suggesting that the polymer can reduce oxidative damage. Interestingly, this protective effect is not seen for synthetic model dopamine pigment where the polymerization process may be incomplete [31]. A further role for neuromelanin appears to be binding a variety of potentially harmful exogenous compounds, such as pesticides and toxins [32–34]. In particular, its role as a metal binder has received attention. Neuromelanin binds a range of metals, including a range of potentially toxic cations [13, 35] and the concentrations of metals associated with the polymer increases with aging [13]. We have demonstrated high- and low-afﬁnity binding sites for iron in neuromelanin [36], suggesting that the interaction of this physiologically important metal with neuromelanin is not random, but highly regulated. The substantia nigra is naturally rich in iron and, given the absence of the iron-binding protein ferritin in the dopaminergic neurons of the human substantia nigra, it is feasible that neuromelanin plays this role as an iron binder within these pigmented cells [37]. Supporting this notion, Mössbauer studies demonstrate many similarities between the physical properties of the iron core in ferritin and the iron clusters found in neuromelanin [38, 39]. Like the iron core in ferritin, neuromelanin binds ferric iron in oxyhydroxy clusters, although these clusters are smaller and less regular than that in ferritin [36]. Proteomic analysis of neuromelanin isolated from the human substantia nigra recently identiﬁed ferritin localized within these granules [40]. It has been suggested that the iron-binding capacity of neuromelanin in vivo is unsaturated [27], thus the pigment may maintain the capacity to bind iron. It is therefore feasible that the pigmented neurons employ this capacity to prevent iron-mediated free radical production. Experimental data using synthetic dopamine melanin (DA-M) supports this hypothesis in that under conditions of low iron, most iron ions are bound to the polymer and free radical production is low [41]. Such data support a protective role for neuromelanin in the healthy human brain. A further biological role of neuromelanin in the degradation pathway of proteins has been suggested from proteomic studies of neuromelanin granules isolated from human substantia nigra [5, 25]. These studies identiﬁed the lysosome integral membrane protein II, cathepsin B and D, and tripeptidyl-peptidase, proteins associated with lysosomes and lysosome-related organelles. It can therefore be hypothesized that, like lysosomes, neuromelanin granules may also play a role in degrading aggregated or misfolded proteins.

8.6 Is Neuromelanin Involved in Neurological Disease?

Despite some evidence for an active and protective role for neuromelanin in the normal brain, the pigment has also been linked with the etiology of the common

233

234

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease a)

b)

2cm c)

d)

1mm Figure 8.5 Neuromelanin pigmentation of the healthy human substantia nigra (a) and the characteristic pallor of this region in the Parkinson’s disease brain seen macroscopically (b). Loss of pigmented cells in the Parkinson’s disease substantia nigra at the microscopic level can be seen in (d),

compared with the healthy substantia nigra in (c). Associated with the death of many of the pigmented neurons in this region is the formation of Lewy bodies (insert in c) in some surviving neurons. (Figure provided with kind permission from Professor Glenda Halliday, Neuroscience Research Australia.)

neurodegenerative disorder Parkinson’s disease. Parkinson’s disease is characterized pathologically by the relatively selective and massive death of the pigmented neurons of the substantia nigra, giving this region a characteristic pale appearance, although the volume of this region is unchanged (Figure 8.5). Correlative analyses of the relative vulnerability of substantia nigra neurons in Parkinson’s disease compared with other pigmented regions of the human brain demonstrated a positive association between regional degree of pigmentation and cell loss in this disorder [42, 43]. Quantitative analyses of pigment volume and the relative vulnerability of pigmented neurons within the various nuclei that make up the substantia nigra, however, does not support the hypothesis that cell vulnerability in Parkinson’s disease is directly related to degree of pigmentation as the differentially vulnerable tiers of this nuclei contain that same amount of pigment [2]. This does not exclude the possibility of a role for the pigment in Parkinson’s

8.7 Effects of Neuromelanin In Vitro and In Vivo

disease. Given the lack of neuromelanin in most laboratory species and the small amount of the pigment in the human brain, particularly in Parkinson’s disease where the amount of pigment is dramatically reduced due to the death of these cells [10, 43], experimental investigations into this question in the human brain are limited. The small amount of available data, however, suggests that changes in the pigment occur in Parkinson’s disease. Pigmented neurons in the parkinsonian brain are reported to contain less pigment than those in the healthy brain [43, 44], but the optical density of the pigment is increased [2]. Biophysical analyzes of neuromelanin isolated from the Parkinson’s disease brain appears to contain a protease-resistant, lipoproteic material not seen in the healthy brain [30, 44] and pigment-associated cholesterol is also reduced [2]. α-Synuclein, a synaptic protein which forms Lewy bodies, abnormal inclusions found in the parkinsonian brain, is cross-linked to neuromelanin pigment from the parkinsonian brain [45]. Further, this protein aggregates speciﬁcally on the vulnerable pigmented cells of the substantia nigra in early disease, suggesting the pigment plays a role in neurodegenerative cascades in this disorder [2]. It has also been reported that the ability of neuromelanin to bind iron is reduced in Parkinson’s disease [46, 47]. This may be signiﬁcant as iron levels have been reported to be increased in the substantia nigra in this disorder [48]. If the iron chelation ability of the pigment is reduced this could potentially result in increased levels of intraneuronal free iron and oxidative stress via the stimulation of free radical production, contributing to perhaps many factors that increase the vulnerability of these pigmented neurons [16].

8.7 Effects of Neuromelanin In Vitro and In Vivo

While factors other than melanization inﬂuence the vulnerability of an individual nigral neuron in Parkinson’s disease [16], pigmentation of these neurons nevertheless appears to play a signiﬁcant role in their eventual fate. Thus, an understanding of the inﬂuence of this pigment is important in understanding the pathogenesis and progression of this disorder. In the remaining sections we review in vitro and in vivo studies investigating the functional effects of neuromelanin. 8.7.1 Mechanisms of Neuromelanin Cytotoxicity

In vitro studies suggest that neuromelanin can induce mitochondria-initiated cell death and inhibit the ubiquitin–proteasome system (UPS) [49–52]. In contrast, however, neuromelanin may also protect neurons by scavenging toxic free radicals, reactive ions, and toxic dopamine quinone, a metabolite of dopamine autoxidation, thus functioning as an antioxidant [53, 54]. These contrasting data suggest that neuromelanin can exert either a neuroprotective or neurotoxic inﬂuence, and that data from cellular and animal experiments with puriﬁed neuromelanin must be carefully interpreted with respect to the human brain.

235

236

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease

Table 8.1 Neurotoxicity of isolated neuromelanin in in vivo and in vitro models.

Melanin

Model system

In vivo experiments

Administered in:

Neuromelanin

Cytotoxicity

Mechanism of cytotoxicity

References

rat substantia nigra

cell loss of dopamine neurons

inﬂammation

[55]

rat substantia nigra (iron-laden neuromelanin)

loss of dopamine

iron-induced oxidative stress

[56]

rat substantia nigra

cell loss of dopamine neurons

glia-mediated inﬂammation

[57]

SH-SY5Y cells

cell death

activation of apoptotic signal

[52, 58]

SH-SY5Y cells

iron-dependent cell death

oxidative stress, UPS inhibition

[49–51]

SK-N-SH, rat mesencephalic cells

no toxicity

proinﬂammatory signaling

[31]

rat microglia

activation

oxidative stress

[59]

mesencephalic neurons

cell death

[60]

rat mesencephalic neurons

no toxicity

[61]

In vitro experiments Neuromelanin

A limited number of in vivo and in vitro experiments investigating the possible cytotoxicity of neuromelanin have been reported, and are summarized in Table 8.1. The data are contradictory, dependent on the model system, amount and type of neuromelanin used, and the measured endpoints. Studies injecting ironsaturated neuromelanin into the rat substantia nigra report a loss of 50% of dopamine neurons compared with controls – a smaller decrease in neuronal number than the 95% loss following injection of free Fe(III) alone [56, 62, 63]. In another study, injection of human neuromelanin into the rat substantia nigra and cerebral cortex induced a signiﬁcant reduction of dopamine cells in the substantia nigra – an effect attributed to strong microglia activation [57]. Activation of microglia by neuromelanin has also been demonstrated in rat primary cultured microglial cells – an effect suggested to be mediated by p38-activated protein kinases [59]. These data may be pertinent for the marked microglial reaction associated with extraneuronal neuromelanin pigment deposits in the human parkinsonian

8.7 Effects of Neuromelanin In Vitro and In Vivo

substantia nigra. In in vitro systems, neuromelanin-mediated cell death appears to be dependent on cell type, as phagocytosis of both iron-saturated neuromelanin and synthetic DA-M stimulates signiﬁcant cell death of neuron-like SK-N-SH cells, but not of glial-like U373 cells [31]. In contrast, phagocytosis of iron-depleted neuromelanin into SK-N-SH cells was not associated with cell death; indeed cell survival following an oxidative stimulus was increased [31]. These data suggest a neuroprotective function of neuromelanin via scavenging cytotoxic iron and reactive oxygen species (ROS) and reactive nitrogen species (RNS) under certain cellular conditions. In contrast, in human dopaminergic SH-SY5Y cells neuromelanin induces mitochondria-dependent apoptosis [52, 64]. Neuromelanin causes mitochondrial membrane permeabilization, permitting transmembrane movement of molecules up to around 1.5 kDa, release of cytochrome c into the cytosol, and activation of an apoptotic cascade, as shown in Figure 8.6. Interestingly, protease K treatment completely prevents induction of this apoptosis, and the cytotoxicity of synthetic DA-M and l-cysteinyl-DA-M (CysDA-M) is much lower than that of human neuromelanin, suggesting the protein component of neuromelanin may play a role in induction of apoptosis. 8.7.2 Neuromelanin Effects on Mitochondrial Function

In Parkinson’s disease, a defect of complex I (NADH : ubiquinone oxidoreductase, EC 1.6.5.3) in the mitochondrial respiratory chain has been reported [66–68]. Complex I is a macromolecule composed of 46 subunits, the activity of which is reduced by impaired subunit assembly. Neuromelanin disturbs the assembly of complex I, but protease K-treated neuromelanin or DA-M does not [52, 58]. Neuromelanin has accessible sulfhydryl groups within its proteinaceous component, the number of which is reduced by protease K treatment to one quarter, suggesting that peptide-bound sulfhydryl residues account for neuromelanin-induced mitochondrial dysfunction and apoptosis. Under physiological conditions or mild oxidative stress, ROS and RNS reversibly modify thiols in neuromelanin, protein, and reduced glutathione (GSH) into active intermediates, thiol radicals, sulfenic (RSOH), and sulﬁnic acid (RSO2H). Activated sulfhydryl groups in protein form mixed disulﬁde (S–S) bonds with those of GSH and other compounds in a reaction called “S-glutathionylation” (Pr-S-SG). Studies using synchrotron X-ray microscopy of human brain tissues have detect these oxidized reactive sulfur compounds in neuromelanin ex vivo [13]. Prolonged or intense exposure to ROS/RNS irreversibly oxidizes protein thiols into cysteine sulfonic acid. Sulfonate has been reported in neuromelanin in the healthy brain [69, 70], and also in the brains of patients with Parkinson’s disease and Alzheimer’s disease [4]. S-Glutathionylation is reversed by glutaredoxin, other thioredoxin, and protein disulﬁde isomerase (EC 5.3.4.1), yielding free protein sulfhydryl and GSH from Pr-S-SG. This reaction is recycled by thioredoxin reductase (EC 1.6.4.5) or GSH reductase (EC 1.6.4.2) using NADPH as a cofactor.

237

238

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease Activation by ROS-RNA NM-SH

a) Deglutathionylation of Complex I

Pr-SSG GSH

Pr-S-SG

Complex I

I II III IV V

I II III IV V

Pr-SH

Dissociation of Complex I, III

Inhibition of ATP synthesis

b) Inhibition of the UPS

Increased ROS-RNS Accumulation of modified proteins a-Synuclein fibril formation

Mitochondrial membrane permeabilization

Lewy body formation Caspase activation c) Induction of apoptosis Control

Figure 8.6 Mechanisms underlying the neurotoxicity of neuromelanin. In cellular models, neuromelanin deglutathionylates mitochondrial proteins and impairs assembly of complex I (a). Mitochondria were incubated in the presence of 10 μg/ml neuromelanin (III), or protease-K-treated neuromelanin (IV) or DA-M (V), or without melanin (II, control). Lane I represents molecular weight markers. After nonreducing sodium dodecyl sulfate–polyacrylamide gel electrophoresis, protein-bound SH residues were detected according to Brennan et al [65]. Complex I was stained with polyclonal antibody speciﬁc for complex I. Increased ROS/RNS and deglutathionylation in mitochondria-induced methylthiazoltetrazolium, and cytochrome c was released as a

+ NM

+ P-K treated NM

function of reaction time: I, II, III and V prior to, and 1, 2, 4 and 6 h after neuromelanin treatment. Caspase-3 was activated in neuromelanin-treated SH-SY5Y cells, but not in control and protease K-treated cells. UPS activity was simultaneously inhibited directly by the oxidative modiﬁcation and indirectly by mitochondrial dysfunction. Inhibition of the 26S proteasome was visualized by accumulation of Green Fluorescent Protein in SH-SY5Y cells transfected with a proteasome sensor vector, pZsProSensor-1 (BD Biosciences) (b). Neuromelanin, but not protease K-treated neuromelanin, induced apoptosis, as shown by Hoechst 33342 cell staining (c). NM, neuromelanin; P-K, protease K.

S-Glutathionylation regulates cellular functions related to energy metabolism, cytoskeletons, signaling-modifying proteins (kinases, phosphatases), Ca2+ homeostasis, protein folding, and apoptosis cascades [71, 72]. S-Glutathionylation stabilizes the assembly of complex I and protects the 4Fe–4S cluster of the 51-kDa subunit [72]. A component of the mitochondrial permeability transition pore,

8.7 Effects of Neuromelanin In Vitro and In Vivo

adenine nucleotide transporter, is S-glutathionylated at Cys57 and the oxidation of this sulfhydryl group with nitric oxide induces mitochondrial membrane permeabilization [73]. In conclusion, glutathionylation appears to be an important mechanism related to neuromelanin function [52, 58]. 8.7.3 Neuromelanin Effects on the UPS

Lewy bodies and Lewy neurites have been suggested to contribute mechanistically to neuronal degeneration in Parkinson’s disease and other synucleinopathy disorders [74, 75], although more recently these inclusions have been considered to be protective [76]. Oxidatively modiﬁed proteins are conjugated with multiple ubiquitin molecules and the tagged protein is degraded by the 26S proteasome complex [77]. UPS dysfunction has been implicated in both familial [78] and sporadic [79] Parkinson’s disease. Interestingly, neuromelanin inhibits the in situ activity of the 26S proteasome [49], increases ROS/RNS production in mitochondria, reduces mitochondrial ATP synthesis, and inhibits the ATP-dependent function of UPS [80, 81]. Increased ROS/RNS also facilitates modiﬁcation of proteins, resulting in proteins that can also inhibit the activity of the UPS [49, 51]. 8.7.4 Comparison of the Cytotoxicity of Neuromelanin with Synthetic DA-M

Synthetic DA-M and CysDA-M have both been employed as models of native neuromelanin, but their composition and biological functions are quite different. Proteins and lipids form major components of the structure of neuromelanin [5, 19, 23, 24], while the melanin component of the pigment is a mixture of indolebased eumelanin and benzothiazine-based pheomelanin, with different oxidation potentials [82, 83]. As summarized in Table 8.2, DA-M consistently induces cell death in animal and cellular models [52, 58, 60, 84–86], primarily via induction of “apoptosis-like” cell death in which ROS/RNS appear to play a decisive role. As summarized in Table 8.3, we have recently studied the cellular mechanisms of apoptosis induced by DA-M compared with that reported for neuromelanin [58]. DA-M oxidatively decreases GSH and sulfhydryl content in mitochondria, whereas neuromelanin increases GSH by dissociation of mixed disulﬁde bonds in complex I. Neuromelanin and DA-M induce mitochondrial membrane permeabilization, and cytochrome c release, whereas Cys-DAM does not. DA-M does not induce signiﬁcant apoptosis, but thiol-targeting reducing reagents, such as GSH, dithiothreitol, and N-acetyl-cysteine, markedly enhance the cytotoxicity. In contrast, neuromelanin directly activates a mitochondria-initiated apoptotic cascade, which is suppressed completely by GSH. In DA-M-treated cells, reducing sulfhydryl reagents activate caspase-3, as shown by detection of activated caspase-3 with a molecular mass of 18 kDa. These results suggest that the pathways leading to cell death initiated by neuromelanin and DA-M differ, depending on

239

240

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease Table 8.2 Neurotoxicity of synthetic DA-M in in vitro models.

Cell system

Cytotoxicity

Mechanism of cytotoxicity

References

SK-N-SH, rat mesencephalic cells

cell death

ROS-mediated apoptosis?

[31]

Rat mesencephalic neurons

cell death

oxidative stress

[52]

SH-SY5Y cells

GSH-dependent apoptosis

mitochondrial membrane permeabilization, GSH-dependent caspase activation

[64]

Rat substantia nigra coculture

cell death

iron-induced oxidative stress

[84]

PC12 cells

apoptosis-like cell death

iron?

[85]

Mouse mesencephalic dopamine cells

apoptosis

[86]

Table 8.3 Comparison of the effects of neuromelanin and DA-M in vitro and in vivo.

Function

Neuromelanin

DA-M

Induction of apoptosis

yes

no; requires reducing sulfhydryl

Activation of apoptotic cascade in mitochondria

Reference

[52, 58, 64]

Induction of mitochondrial membrane permeabilization

yes

yes

Release of cytochrome c

yes

yes

Reduction of membrane potential (DYm)

yes

yes

Reduced ATP synthesis

yes

yes

Increased ROS/RNS production

yes

yes

Caspase-3 activation

yes

no; requires GSH

Protection by Bcl-2

yes

no

S-Glutathionylation, S-deglutathionylation

in mitochondria

no

Redox state in mitochondria

to reduced state

to oxidized state

Redox state in cytoplasm

no signiﬁcant effect

to oxidative state

inhibition of 26S

no signiﬁcant effect

[52, 64]

Effects on redox state

Effects on the UPS

[52, 58]

[49–51]

References

the differential regulation of the redox state in the cytoplasm and mitochondria [58, 64]. In summary, neuromelanin primarily perturbs S-glutathionylation in mitochondria and inhibits this function as the initial event in the cell death pathway, whereas DA-M oxidatively modiﬁes proteins in subcellular components, especially mitochondria and the UPS. These results strongly suggest cytotoxicity mediated by neuromelanin is attributable not only to its melanin composition, but also to the proteinaceous and other components of this molecule.

8.8 Conclusions

Despite the advances of recent years, neuromelanins represent an enigmatic member of the pigment family, about which much still remains to be discovered. Nevertheless, it is clear that these pigments are not simply an inert waste product of neuronal metabolism as previously believed, but play important roles in neuronal function in the human brain. Neuromelanin appears to protect neurons under physiological conditions, but it may also promote cellular toxicity in certain degenerative conditions. Speciﬁcally, the neuromelanin of the substantia nigra appears to be involved in a number of pathways accepted to be involved in neurodegenerative cascades in Parkinson’s disease and related disorders. Characterization of the role of neuromelanin in the healthy and diseased brain will improve our understanding of this interesting pigment and our understanding of the etiology of a number of common neurodegenerative disorders.

Acknowledgments

K.L.D. holds a Research Fellowship from the National Health and Medical Research Council of Australia. This work was supported by the National Health and Medical Research Council of Australia, and a Grant-in-Aid for Scientiﬁc Research (C) of the Japanese Ministry of Education, Culture and Science and a Grant-in-Aid for Clinical Research Promotion of the Japanese Ministry of Welfare and Health (W. M.). K.L.D. and W.M. contributed equally to this work.

References 1 Elleder, M. and Borovansky, J. (2001)

Autoﬂuorescence of melanins induced by ultraviolet radiation and near ultraviolet light. A histochemical and biochemical study. Histochem. J., 33, 273–281. 2 Halliday, G.M., Ophof, A., Broe, M., Jensen, P.H., Kettle, E., Fedorow, H., Cartwright, M., Grifﬁths, F.M., Shepherd, C.E., and Double, K.L. (2005) α-Synuclein

redistributes to neuromelanin lipid in the substantia nigra early in Parkinson’s disease. Brain, 128, 2654–2664. 3 Double, K.L., Dedov, V.N., Fedorow, H., Kettle, E., Halliday, G., Garner, B., and Brunk, U.T. (2008) The comparative biology of neuromelanin and lipofuscin in the human brain. Cell. Mol. Life Sci., 65, 1669–1682.

241

242

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease 4 Sulzer, D., Bogulavsky, J., Larsen, K.,

5

6

7

8

9

10

11

12

Behr, G., Karatekin, E., Kleinman, M., Turro, N., Krantz, D., Edwards, R., Greene, L., et al. (2000) Neuromelanin biosynthesis is driven by excess cytosolic catecholamines not accumulated by synaptic vesicles. Proc. Natl. Acad. Sci. USA, 97, 11869–11874. Tribl, F., Gerlach, M., Marcus, K., Asan, E., Tatschner, T., Arzberger, T., Meyer, H.E., Bringmann, G., and Riederer, P. (2005) “Subcellular proteomics” of neuromelanin granules isolated from the human brain. Mol. Cell Proteomics, 4, 945–947. Bogerts, B. (1981) A brainstem atlas of catecholaminergic neurons in man, using melanin as a natural marker. J Comp. Neurol., 197, 63–80. Saper, C.B. and Petito, C.K. (1982) Correspondence of melanin-pigmented neurons in human brain with A1–A14 catecholamine cell groups. Brain, 105, 87–101. Zecca, L., Bellei, C., Costi, P., Albertini, A., Monzani, E., Casella, L., Gallorini, M., Bergamaschi, L., Moscatelli, A., Turro, N.J., et al. (2008) New melanic pigments in the human brain that accumulate in aging and block environmental toxic metals. Proc. Natl. Acad. Sci. USA, 105, 17567–17572. Fedorow, H., Halliday, G.M., Rickert, C., Gerlach, M., Riederer, P., and Double, K.L. (2006) Evidence for speciﬁc phases in the development of human neuromelanin. Neurobiol. Aging, 27, 506–512. Zecca, L., Fariello, R., Riederer, P., Sulzer, D., Gatti, A., and Tampellini, D. (2002) The absolute concentration of nigral neuromelanin, assayed by a new sensitive method, increases throughout the life and is dramatically decreased in Parkinson’s disease. FEBS Lett., 510, 216–220. Tribl, F., Arzberger, T., Riederer, P., and Gerlach, M. (2007) Tyrosinase is not detected in human catecholaminergic neurons by immunohistochemistry and Western blot analysis. J. Neural Transm. Suppl., (72), 51–55. Check, E. (2002) Parkinson’s disease patients show positive response to implants. Nature, 416, 666.

13 Bohic, S., Murphy, K., Paulus, W.,

14

15

16

17

18

19

20

21

Cloetens, P., Salomé, M., Susini, J., and Double, K. (2008) Intracellular chemical imaging of the developmental phases of human neuromelanin using synchrotron X-ray microspectroscopy. Anal. Chem., 80, 9557–9566. Forno, L.S. (1996) Neuropathology of Parkinson’s disease. J. Neuropathol. Exp. Neurol., 55, 259–272. Langston, J.W., Forno, L.S., Tetrud, J., Reeves, A.G., Kaplan, J.A., and Karluk, D. (1999) Evidence of active nerve cell degeneration in the substantia nigra of humans years after 1-methyl-4-phenyl1,2,3,6-tetrahydropyridine exposure. Ann. Neurol., 46, 598–605. Double, K.L., Reyes, S., Werry, E.L., and Halliday, G.M. (2010) Selective cell death in neurodegeneration: why are some neurons spared in vulnerable regions? Prog. Neurobiol., 92, 316–329. Beach, T.G., Sue, L.I., Walker, D.G., Lue, L.F., Connor, D.J., Caviness, J.N., Sabbagh, M.N., and Adler, C.H. (2007) Marked microglial reaction in normal aging human substantia nigra: correlation with extraneuronal neuromelanin pigment deposits. Acta Neuropathol., 114, 419–424. Fedorow, H., Tribl, F., Halliday, G., Gerlach, M., Riederer, P., and Double, K.L. (2005) Neuromelanin in human dopamine neurons: comparison with peripheral melanins and relevance to Parkinson’s disease. Prog. Neurobiol., 75, 109–124. Double, K., Zecca, L., Costo, P., Mauer, M., Greisinger, C., Ito, S., Ben-Shachar, D., Bringmann, G., Fariello, R.G., Riederer, P., et al. (2000) Structural characteristics of human substantia nigra neuromelanin and synthetic dopamine melanins. J. Neurochem., 75, 2583–2589. Ichinose, H., Ohye, T., Fujita, K., Pantucek, F., Lange, K., Riederer, P., and Nagatsu, T. (1994) Quantiﬁcation of mRNA of tyrosine hydroxylase and aromatic l-amino acid decarboxylase in the substantia nigra in Parkinson’s disease and schizophrenia. J. Neural Transm., 8, 149–158. Lehmann, I.T., Bobrovskaya, L., Gordon, S.L., Dunkley, P.R., and Dickson, P.W.

References

22

23

24

25

26

27

28

29

30

(2006) Differential regulation of the human tyrosine hydroxylase isoforms via hierarchical phosphorylation. Biol. Chem., 281, 17644–17651. Fedorow, H., Pickford, R., Hook, J.M., Double, K.L., Halliday, G.M., Gerlach, M., Riederer, P., and Garner, B. (2005) Dolichol is the major lipid component of human substantia nigra neuromelanin. J. Neurochem., 92, 990–995. Dzierzega-Lecznar, A., Kurkiewicz, S., Stepien, K., Chodurek, E., Riederer, P., and Gerlach, M. (2006) Structural investigations of neuromelanin by pyrolysis-gas chromatography/mass spectrometry. J. Neural Transm., 113, 729–734. Zecca, L., Costi, P., Mecacci, C., Ito, S., Terreni, M., and Sonnino, S. (2000) Interaction of human substantia nigra neuromelanin with lipids and peptides. J. Neurochem., 74, 1758–1765. Tribl, F., Marcus, K., Meyer, H.E., Bringmann, G., Gerlach, M., and Riederer, P. (2006) Subcellular proteomics reveals neuromelanin granules to be a lysosome-related organelle. J. Neural Transm., 113, 741–749. Shen, X., Zhang, F., and Dryhurst, G. (1997) Oxidation of dopamine in the presence of cysteine: characterization of new toxic products. Chem. Res. Toxicol., 10, 147–155. Shima, T., Sarna, T., Swartz, H., Stroppolo, A., Gerbasi, R., and Zecca, L. (1997) Binding of iron to neuromelanin of human substantia nigra and synthetic melanin: an electron paramagnetic resonance spectroscopy study. Free Radic. Biol. Med., 23, 110–119. Zecca, L., Zucca, F.A., Wilms, H., and Sulzer, D. (2003) Neuromelanin of the substantia nigra: a neuronal block hole with protective and toxic characteristics. Trends Neurosci., 26, 578–580. Rozanowska, M., Sarna, T., Land, E., and Truscott, T. (1999) Free radical scavenging properties of melanin interaction of eu- and pheo-melanin models with reducing and oxidising radicals. Free Radical. Biol. Med., 26, 518–525. Fasano, M., Bergamasco, B., and Lopiano, L. (2006) Is neuromelanin

31

32

33

34

35

36

37

38

39

changed in Parkinson’s disease? Investigations by magnetic spectroscopies. J. Neural Transm., 113, 769–774. Li, J., Scheller, C., Koutsilieri, E., Grifﬁths, F., Beart, P.M., Mercer, L.D., Halliday, G., Kettle, E., Rowe, D., Riederer, P., et al. (2005) Differential effects of human neuromelanin and synthetic dopamine melanin on neuronal and glial cells. J. Neurochem., 95, 599–608. D’Amato, R.J., Lipman, Z.P., and Snyder, S.H. (1986) Selectivity of the parkinsonian neurotoxin MPTP: toxic metabolite MPP+ binds to neuromelanin. Science, 231, 987–989. Lindquist, N.G., Larsson, B.S., and Lyden-Sokolowski, A. (1988) Autoradiography of (14C)paraquat or (14C) diquat in frogs and mice: accumulation in neuromelanin. Neurosci Lett., 93, 1–6. Ostergren, A., Annas, A., Skog, K., Lindquist, N.G., and Brittlebo, E.B. (2004) Long-term retention of neurotoxic beta-carbolines in brain neuromelanin. J. Neural Transm., 111, 141–157. Zecca, L., Tampellini, D., Gatti, A., Crippa, R., Eisner, M., Sulzer, D., Ito, S., Fariello, R., and Gallorini, M. (2002) The neuromelanin of the human substantia nigra and its interaction with metals. J. Neural Transm., 109, 663–672. Double, K.L., Gerlach, M., Schünemann, V., Trautwein, A.X., Zecca, L., Gallorini, M., Youdim, M.B.H., Riederer, P., and Ben-Shachar, D. (2003) Iron binding characteristics of neuromelanin of the human substantia nigra. Biochem. Pharmacol., 66, 489–494. Ben-Shachar, D., Riederer, P., and Youdim, M.B.H. (1991) Iron–melanin interaction and lipid peroxidation: implications for Parkinson’s disease. J. Neurochem., 57, 1609–1614. Gerlach, M., Trautwein, A.X., Zecca, L., Youdim, M.B.H., and Riederer, P. (1995) Mössbauer spectroscopic studies of puriﬁed human neuromelanin isolated from the substantia nigra. J. Neurochem., 65, 923–926. Galazka-Friedman, J., Bauminger, E.R., Friedman, A., Barcikowska, M., Hechel, D., and Nowik, I. (1996) Iron in

243

244

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease

40

41

42

43

44

45

46

47

parkinsonian and control substantia nigra – a Mossbauer spectroscopy study. Mov. Disord., 11, 8–16. Tribl, F., Asan, E., Arzberger, T., Tatschner, T., Langenfeld, E., Meyer, H.E., Bringmann, G., Riederer, P., Gerlach, M., and Marcus, K. (2009) Identiﬁcation of l-ferritin in neuromelanin granules of the human substantia nigra: a targeted proteomics approach. Mol. Cell Proteomics, 8, 1832–1838. Zareba, M., Bober, A., Korytowski, W., Zecca, L., and Sarna, T. (1995) The effect of a synthetic neuromelanin on yield of free hydroxyl radicals generated in model systems. Biochem. Biophys. Acta, 1271, 343–348. Hirsch, E., Graybiel, A., and Agid, Y. (1988) Melanized dopamine neurons are differentially susceptible to degeneration in Parkinson’s disease. Nature, 28, 345–348. Kastner, A., Hirsch, E., Lejeune, O., Javoy-Agid, F., Rascol, O., and Agid, Y. (1992) Is the vulnerability of neurons in the substantia nigra of patients with Parkinson’s disease related to their neuromelanin content? J. Neurochem., 59, 1080–1089. Aime, S., Bergamasco, B., Casu, M., Digilio, G., Fasano, M., Giraudo, S., and Lopiano, L. (2000) Isolation and 13C-NMR characterization of an insoluble proteinaceous fraction from substantia nigra of patients with Parkinson’s disease. Mov. Disord., 15, 977–981. Fasano, M., Giraudo, S., Coha, S., Bergamasco, B., and Lopiano, L. (2003) Residual substantia nigra neuromelanin in Parkinson’s disease is cross-linked to alpha-synuclein. Neurochem. Int., 42, 603–606. Bolzoni, F., Giraudo, S., Lopiano, L., Bergamasco, B., Fasano, M., and Crippa, P.R. (2002) Magnetic investigations of human mesencephalic neuromelanin. Biochem. Biophys. Acta, 1586, 210–218. Lopiano, L., Chiesa, M., Digilio, D., Giraudo, G., Bergamasco, B., and Fasano, M. (2000) Q-band EPR investigations of neuromelanin in control and Parkinson’s disease patients. Biochem. Biophys. Acta, 1500, 306–312.

48 Lee, D.W. and Andersen, J.K. (2010) Iron

49

50

51

52

53

54

55

elevations in the aging Parkinsonian brain: a consequence of impaired iron homeostasis? J. Neurochem., 112, 332–339. Shamoto-Nagai, M., Maruyama, W., Akao, Y., Osawa, T., Tribl, F., Gerlach, M., Zucca, F.A., Zecca, L., Riederer, P., and Naoi, M. (2004) Neuromelanin inhibits enzymatic activity of 26S proteasome in human dopaminergic SH-SY5Y cells. J. Neural Transm., 111, 1253–1265. Shamoto-Nagai, M., Maruyama, W., Yi, H., Akao, Y., Tribl, F., Gerlach, M., Osawa, T., Riederer, P., and Naoi, M. (2006) Neuromelanin induces oxidative stress in mitochondria through release of iron: mechanism behind the inhibition of 26S proteasome. J. Neural Transm., 113, 633–644. Maruyama, W., Shamoto-Nagai, M., Akao, Y., Riederer, P., and Naoi, M. (2006) The effect of neuromelanin on the proteasome activity in human dopaminergic SH-SY5Y cells. J. Neural Transm. Suppl., 70, 125–132. Naoi, M., Maruyama, W., Yi, H., Yamaoka, Y., Shamoto-Nagai, M., Akao, Y., Gerlach, M., Tanaka, M., and Riederer, P. (2008) Neuromelanin selectively induces apoptosis in dopaminergic SH-SY5Y cells by deglutathionylation in mitochondria: involvement of the protein and melanin component. J. Neurochem., 105, 2489–2500. Double, K.L., Ben-Shachar, D., Youdim, M.B., Zecca, L., Riederer, P., and Gerlach, M. (2002) Inﬂuence of neuromelanin on oxidative pathways within the human substantia nigra. Neurotoxicol. Teratol., 24, 621–628. Zecca, L., Casella, L., Albertini, A., Bellei, C., Zucca, F.A., Engelen, M., Zadlo, A., Szewczyk, G., Zareba, M., and Sarna, T. (2008) Neuromelanin can protect against iron-mediated oxidative damage in system modeling iron overload of brain aging and Parkinson’s disease. J. Neurochem., 106, 1866–1875. Gerlach, M. and Riederer, P. (1996) Animal models of Parkinson’s disease: an empirical comparison with the

References

56

57

58

59

60

61

62

63

phenomenology of the disease in man. J. Neural Transm., 103, 987–1041. Double, K.L., Halliday, G.M., Henderson, J., Grifﬁths, F.M., Heinemann, T., Riederer, P., and Gerlach, M. (2003) The dopamine receptor agonist lisuride attenuates iron-mediated dopaminergic neurodegeneration. Exp. Neurol., 184, 530–535. Zecca, L., Wilms, H., Geick, S., Claasen, J.H., Brandenburg, L.O., Holzknecht, C., Panizza, M.L., Zucca, F.A., Deuschl, G., Sievers, J., et al. (2008) Human neuromelanin induces neuroinﬂammation and neurodegeneration in the rat substantia nigra: implications for Parkinson’s disease. Acta Neuropathol., 116, 47–55. Naoi, M., Maruyama, W., Yi, H., Inaba, K., Akao, Y., and Shamoto-Nagai, M. (2009) Mitochondria in neurodegenerative disorders: regulation of the redox state and death signaling leading to neuronal death and survival. J. Neural. Transm., 116, 1371–1381. Wilms, H., Rosenstiel, P., Sievers, J., Deuschl, G., Zecca, L., and Lucius, R. (2003) Activation of microglia by human neuromelanin is NF-κB-dependent and involves p38 mitogen-activated protein kinase: implications for Parkinson’s disease. FASEB J., 17, 500–502. Depboylu, C., Matusch, A., Tribl, F., Zoriy, M., Michel, P.P., Riederer, P., Gerlach, M., Becker, S., Oertel, W.H., and Hoglinger, G.U. (2007) Glia protects neurons against extracellular human neuromelanin. Neurodegener. Dis., 4, 218–226. Double, K.L. (2006) Functional effects of neuromelanin and synthetic melanin in model systems. J. Neural Transm., 113, 751–756. Gerlach, M., Desser, H., Youdim, M.B.H., and Riederer, P. (1996) New horizons in molecular mechanisms underlying Parkinson’s disease and in our understanding of the neuroprotective effects of selegiline. J. Neural Transm., 48, 7–21. Gerlach, M., Riederer, P., and Double, K.L. (2008) Neuromelanin-bound ferric iron as an experimental model of dopaminergic neurodegeneration in

64

65

66

67

68

69

70

71

Parkinson’s disease. Parkinsonism Relat. Disord., 14 (Suppl. 2), S185–S188. Naoi, M., Yi, H., Maruyama, W., Inaba, K., Shamoto-Nagai, M., Akao, Y., Gerlach, M., and Riederer, P. (2009) Glutathione redox status in mitochondria and cytoplasm differentially and sequentially activates apoptosis cascade in dopaminemelanin-treated SH-SY5Y cells. Neurosci. Lett., 465, 118–122. Brennan, J.P., Wait, R., Begum, S., Bell, J.R., Dunn, M.J., and Eaton, P. (2004) Detection and mapping of widespread intermolecular protein disulﬁde formation during cardiac oxidative stress using proteomics with diagonal electrophoresis. J. Biol. Chem., 279, 41352–41360. Reichmann, H. and Riederer, P. (1989) Biochemische Analyse der Atmungskettenkomplex verschiedener Hirnregionen von Patient mit M. Parkinson und andere Basalganglienerkrankungen. Symposium des BMFT “Morbus Parkinson und andere Basalganglienerkrankungen,” Bad Kissingen. Mizuno, Y., Ohta, S., Tanaka, M., Takamiya, S., Suzuki, K., Sato, T., Oya, H., Ozawa, T., and Kagawa, Y. (1989) Deﬁciencies in complex I subunits of the respiratory chain in Parkinson’s disease. Biochem. Biophys. Res. Commun., 163, 1450–1455. Schapira, A.H., Cooper, J.M., Dexter, D., Jenner, P., Clark, J.B., and Marsden, C.D. (1989) Mitochondrial complex I deﬁciency in Parkinson’s disease. Lancet, 1, 1269. Barden, H. (1984) The oxidative generation of sulfonic acid groups in neuromelanin and lipofuscin in the human brain. J. Histochem. Cytochem., 32, 329–336. Choi, J., Rees, H.D., Weintraub, S.T., Levey, A.I., Chin, L.S., and Li, L. (2005) Oxidative modiﬁcations and aggregation of Cu,Zn-superoxide dismutase associated with Alzheimer and Parkinson diseases. J. Biol. Chem., 280, 11648–11655. Townsend, D.M. (2007) SGlutathionylation: indicator of cell stress and regulator of the unfolded protein response. Mol. Interv., 7, 313–324.

245

246

8 Biological Role of Neuromelanin in the Human Brain and Its Importance in Parkinson’s Disease 72 Mieyal, J.J., Gallogly, M.M., Qanungo, S.,

80 Shamoto-Nagai, M., Maruyama, W., Kato,

Sabens, E.A., and Shelton, M.D. (2008) Molecular mechanisms and clinical implications of reversible protein S-glutathionylation. Antioxid. Redox Signal., 10, 1941–1988. Costantini, P., Belzacq, A.S., Vieira, H.L., Larochette, N., de Pablo, M.A., Zamzami, N., Susin, S.A., Brenner, C., and Kroemer, G. (2000) Oxidation of a critical thiol residue of the adenine nucleotide translocator enforces Bcl-2-independent permeability transition pore opening and apoptosis. Oncogene, 19, 307–314. Ross, C.A. and Poirier, M.A. (2004) Protein aggregation and neurodegenerative disease. Nat. Med., 10 (Suppl.), S10–S17. McNaught, K.S., Olanow, C.W., Halliwell, B., Isacson, O., and Jenner, P. (2001) Failure of the ubiquitin–proteasome system in Parkinson’s disease. Nat. Rev. Neurosci., 2, 589–594. Halliday, G.M. and McCann, H. (2008) Human-based studies on alpha-synuclein deposition and relationship to Parkinson’s disease symptoms. Exp. Neurol., 209, 12–21. Glickman, M.H. and Ciechanover, A. (2002) The ubiquitin–proteasome proteolytic pathway: destruction for the sake of construction. Physiol. Rev., 82, 373–428. Cook, C. and Petrucelli, L. (2009) A critical evaluation of the ubiquitin– proteasome system in Parkinson’s disease. Biochim. Biophys. Acta, 1792, 664–675. McNaught, K.S., Belizaire, R., Isacson, O., Jenner, P., and Olanow, C.W. (2003) Altered proteasomal function in sporadic Parkinson’s disease. Exp. Neurol., 179, 38–46.

Y., Isobe, K., Tanaka, M., Naoi, M., and Osawa, T. (2003) An inhibitor of mitochondrial complex I, rotenone, inactivates proteasome by oxidative modiﬁcation and induces aggregation of oxidized proteins in SH-SY5Y cells. J. Neurosci. Res., 74, 589–597. Sawada, H., Kohno, R., Kihara, T., Izumi, Y., Sakka, N., Ibi, M., Nakanishi, M., Nakamizo, T., Yamakawa, K., Shibasaki, H., et al. (2004) Proteasome mediates dopaminergic neuronal degeneration, and its inhibition causes alpha-synuclein inclusions. J. Biol. Chem., 279, 10710–10719. Odh, G., Carstam, R., Paulson, J., Wittbjer, A., Rosengren, E., and Rorsman, H. (1994) Neuromelanin of the human substantia nigra: a mixed-type melanin. J. Neurochem., 62, 2030–2036. Samokhvalov, A., Hong, L., Liu, Y., Garguilo, J., Nemanich, R.J., Edwards, G.S., and Simon, J.D. (2005) Oxidation potentials of human eumelanosomes and pheomelanosomes. Photochem. Photobiol., 81, 145–148. Mochizuki, H., Nishi, K., and Mizuno, Y. (1993) Iron–melanin complex is toxic to dopaminergic neurons in nigrostriatal co-culture. Neurodegeneration, 2, 1–7. Offen, D., Ziv, I., Barzilai, A., Gorodin, S., Glater, E., Hochman, A., and Melamed, E. (1997) Dopamine-melanin induces apoptosis in PC12 cells: possible implications for the etiology of Parkinson’s disease. Neurochem. Int., 31, 207–216. Nguyen, A., Gille, G., Moldzio, R., Hung, S.-T., and Rausch, W.-D. (2002) Synthetic neuromelanin is toxic to dopaminergic cell cultures. J. Neural Transm., 109, 651–661.

73

74

75

76

77

78

79

81

82

83

84

85

86

247

9 Biogenesis of Melanosomes Cédric Delevoye, Francesca Giordano, Michael S. Marks, and Graça Raposo

9.1 Introduction

Melanin pigments are synthesized within epidermal and ocular melanocytes and the retinal pigment epithelium (RPE) in specialized organelles called melanosomes. Synthesis and sequestration of melanin in membrane-enclosed compartments within these cells prevents exposure of most cellular components to oxidative reactants that are produced during synthesis and that would otherwise be deleterious. Melanins produced in melanocytes in the skin are transferred to keratinocytes, providing photoprotection against ionizing UV radiation, but also resulting in the characteristic pigmentation of skin and hair, and the very different visual cues in the animal kingdom [1]. In the eye, melanins synthesized within melanosomes of RPE cells are not released, but rather are retained within these cells where they function in focusing light and in detoxifying free radicals liberated from phagocytosed photoreceptor outer membrane segments [2]. Melanosomes are also found in the eye in melanocytes within the choroid and in the pigmented epithelium of the iris and ciliary body. The timing of melanin synthesis and thereby also of melanosome production differs in skin melanocytes and in the choroid and RPE. Whereas in the skin melanosomes are produced throughout life, in the RPE the majority of melanin synthesis occurs during embryonic and early postnatal life [3]. To accomplish their role as melanin factories within the pigment cell, melanosomes have unique morphological and compositional features. They are membrane-bound organelles that bear melanocyte-speciﬁc proteins involved in melanosome structure, melanin synthesis, and in maintenance of the most favorable ionic, redox, and osmotic environment for pigment synthesis. Melanosomes share a number of properties associated with conventional lysosomes, the major degradative compartment of the cell, and are therefore classiﬁed as members of a family of so-called lysosome-related organelles (LROs). Melanosomes have a low luminal pH, and contain lysosomal hydrolases and lysosomal membrane proteins (reviewed in [4]). Although melanocytes can make both brown/black eumelanin

Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

248

9 Biogenesis of Melanosomes

and yellow/red pheomelanin, melanosomes that make predominantly pheomelanins differ in structure and composition, and much less is known about their biogenesis. For this reason this chapter focuses on melanocytes that make predominantly eumelanosomes. The biogenesis of eumelanosomes is a multistep process in which an immature unpigmented precursor organelle forms and then subsequently matures to a fully pigmented structure. Such maturation requires morphogenetic and structural modiﬁcations of endosomal intermediates accompanied by interorganellar transport of protein cargos. These cargoes largely consist of melanocyte-speciﬁc components required for melanosome structure and pigment synthesis, including melanogenic enzymes, transporters, and structural components. Mutations in the genes encoding many of these components result in oculocutaneous albinism (OCA) or ocular albinism (OA) due to a failure in melanosome maturation/ function. Melanosome maturation is also accompanied by the recruitment of effector proteins necessary for the appropriate transport of melanosomes to the cell periphery (as discussed in Chapter 10). Over the past few years, studies on skin melanocytes and pigmented melanoma cells have started to unravel the major pathways and molecular mechanisms used by melanocytes to generate eumelanosomes. These studies have highlighted how eumelanosomes become segregated from the main “conventional” endolysosomal system [5–7]. Our emerging understanding of melanosome biogenesis serves as an excellent model that likely applies to the formation of other LROs in different cell types. Like all other membrane and lumenal proteins of the endocytic system, melanosomal proteins are synthesized in the endoplasmic reticulum (ER) and processed during secretory trafﬁc through the Golgi complex. From the Golgi, these cargoes are targeted to melanosomes via specialized post-Golgi trafﬁcking routes that converge upon the endosomal system. The endosomal pathways through which these proteins trafﬁc are only beginning to be understood, and are regulated by both ubiquitous and cell-type-speciﬁc effectors. Ubiquitous trafﬁcking effectors include the heterotetrameric adaptors that recognize speciﬁc cytoplasmic targeting signals on cargo proteins, small GTPases of the Rab superfamily that recruit effectors from the cytoplasm to endosomal intermediates, SNARE (soluble Nethylmaleimide-sensitive factor attachment protein receptor) proteins that specify membrane fusion, and cytoskeleton-associated molecular motors (as detailed in Section 9.4). Importantly, the genes encoding some components of these and other requisite molecular machineries bear mutations in syndromic genetic diseases that are characterized in part by hypopigmentation due to melanosome dysfunction. Among these diseases are the Chediak–Higashi syndrome (CHS) and several variants of the Hermansky–Pudlak syndrome (HPS) [8]. A number of ﬁndings over the last 10 years have begun to unravel how pigment cells integrate unique and ubiquitous molecular mechanisms in specializing the endosomal system to generate cell-type-speciﬁc structures and their associated functions. These studies have shed light not only on the biogenesis of melanosomes and other LROs, but also on general aspects of vesicular transport in the endocytic system and major dysfunctions in genetic diseases.

9.2 Melanosomes: Intracellular Organelles Specialized in Melanin Synthesis

9.2 Melanosomes: Intracellular Organelles Specialized in Melanin Synthesis 9.2.1 Melanosomes Are Unique Organelles That Develop through Different Stages

As melanins are highly electron-dense, melanosomes within epidermal melanocytes were among the ﬁrst organelles to be well characterized by electron microscopy [9]. Eumelanosomes – containing black and brown melanins – can be divided into four stages, termed I–IV, based on their morphology [10] (Figure 9.1). The earliest stages – also often referred to as premelanosomes – lack pigment, but are characterized by the formation of a ﬁbrillar matrix upon which melanins are deposited in later stages. Stage I melanosomes, which are accessible to endocytic tracers and correspond to an early endosomal compartment [5], contain a few

Ultrastructure of melanosomes. Electron microscopy of MNT-1 human melanoma cells ﬁxed by high-pressure freezing before cryosubstitution and embedding in Epon. The four stages of melanosome development are shown in the lower panels. Note the dense bilayered coat (arrowhead) and intralumenal vesicles

Figure 9.1

(arrow) of stage I melanosomes, the proteinaceous ﬁbrils (arrow) of stage II, and the melanin deposition (black) in stages III and IV. The main panel shows a typical ﬁeld of MNT-1 cytoplasm near the nucleus, which contains all four stages of melanosomes. m, mitochondria; N, nucleus; GA, Golgi apparatus. Scale bars: 200 nm.

249

250

9 Biogenesis of Melanosomes

irregular ﬁbrils and intralumenal membrane vesicles. Interestingly, the ﬁbrils emanate from the intralumenal membranes and elongate as the organelle matures [11]. Stage II melanosomes are characterized by fully matured parallel arrays of ﬁbrils organized into concentric sheets [10, 11]. Upon initiation of melanin synthesis, melanin polymers deposit on these ﬁbrillar sheets, resulting in their thickening and blackening as observed by electron microscopy [5, 10]. Continued melanin synthesis results in melanin deposition throughout the melanosome and masking of the underlying contents. Pigmented eumelanosomes in which ﬁbrillar sheets are still apparent are referred to as stage III melanosomes, whereas those in which the underlying ﬁbrillar matrix is masked are considered stage IV [10]. In cells that make predominantly pheomelanins, the melanin content of melanosomes is dark and the underlying ﬁbrillar structure is absent [12]. The formation of melanosomes in eye pigment cells has been much less studied than in skin melanocytes. Genetic analyses in mice with coat-color dilution and in OCA patients (reviewed in [13]), as well as a few limited studies of melanogenesis in early eye development, suggest that the mechanisms underlying melanosome biogenesis in eye pigment cells is similar to that in skin melanocytes. In the RPE and choroid, melanosomes also appear to develop progressively from an immature unpigmented precursor to a mature pigmented melanosome [14]. However, melanogenesis in the RPE occurs predominantly before birth and is largely completed soon after birth, such that there is very little melanin synthesis and melanosome renewal in the adult [15, 16]. Melanogenesis in adult RPE can be induced under certain experimental conditions [17, 18] and can occur in structures without the characteristic striated precursors [19], but it is not clear whether this can occur physiologically. How are the different stage melanosomes structures generated? Both the formation of the underlying ﬁbrillar matrix in stage I and II melanosomes and melanin synthesis/deposition in stage III and IV melanosomes are dependent upon pigment cell-speciﬁc proteins that are speciﬁcally targeted to newly forming organelles. While melanosomes were once considered modiﬁed lysosomes because of their content of lysosomal membrane proteins and enzymes [4, 20–23], they are now appreciated to coexist with lysosomes and other organelles of the late endocytic pathway [5]. As described in detail below, stage I and II melanosomes develop from vacuolar domains of early endosomal compartments that are enriched in the pigment cell-speciﬁc protein, Pmel17 (also known as gp100 or SILV). Current models suggest that stage II melanosomes must be fully “matured” – including fully formed ﬁbrillar sheets – before they become competent to synthesize melanins. Melanin synthesis to generate stage III and IV melanosomes coincides with the delivery to the mature stage II compartments of melanogenic enzymes and transporters that shape the environment of the organelle to favor melanin synthesis (reviewed in [7]). Below we describe these components and our current understanding of these trafﬁcking pathways.

9.2 Melanosomes: Intracellular Organelles Specialized in Melanin Synthesis

9.2.2 Melanosomal Components

Most known melanosomal components are integral membrane proteins that are uniquely expressed in pigment cells. Mutations in the genes that encode many of these proteins cause coat-color dilution in animals or albinism in humans – characterized by variable loss of pigmentation in the skin, hair, and/or eyes due to an impairment of the number, structure, and/or function of melanosomes [24] (Table 9.1). Table 9.1

Melanosome main components and their functions.

Melanosomal proteins

Melanosome stages

Function

Human disease/ mouse model

Tyrosinase

III/IV

melanin synthesis

OCA1 (h)/TyrC (albino)

oxidation of tyrosinase to dopa Tyrp1

III/IV

melanin synthesis oxidation of DHICA to eumelanin

Tyrp2/Dct

III/IV

melanin synthesis tautomerization of dopachrome to DHICA

OCA3 (h)/Tyrp1b (brown) unknown/Dctslt (slaty)

Pmel17/ gp100/ME20

I–II (epitopes masked by melanin in III and IV)

component of the amyloid ﬁbrillar sheets

unknown/si (silver)

P/OCA2

III/IV

melanosome acidiﬁcation

OCA2 (h)/p (pink-eyed dilute)

putative anion transporter MATP/ SLC45A2

unknown

putative membrane transporter

OCA4 (h)

SLC24A5

unknown

putative membrane transporter

unknown/OA

OA1/ GPR143

II–III–IV–lysosomes (bulk is in II)

GPCR

OA1/Oa1

MART-1

I–II

accessory protein for Pmel17 and OA1 function

control of melanosome composition and size unknown

human melanoma antigen OCA1, OCA2, OCA3, OCA4: oculocutaneous albinism type 1, 2, 3, and 4, respectively; OA1: ocular albinism type 1.

251

252

9 Biogenesis of Melanosomes

Pmel17 serves as the structural foundation of the ﬁbrils in stage I and II melanosomes. Pmel17 is synthesized as a transmembrane protein, but is proteolytically processed to smaller lumenal fragments that polymerize to form the intralumenal ﬁbrillar sheets (reviewed in [25]). The mechanisms regulating Pmel17 maturation are discussed below. The accumulation of Pmel17 ﬁbrils is the only currently known deﬁning factor for stage II melanosomes, although one study has suggested that autophagy markers, such as lipidated LC3a, are also present within these organelles [26], suggesting that premelanogenesis may require the same molecular machinery used by the autophagy pathway. Consistent with the onset of melanin synthesis in the transition from stage II melanosomes, the melanin biosynthetic enzymes tyrosinase, tyrosinase-related protein 1 (Tyrp1) and dopachrome tautomerase (Dct; also known as Tyrp2) are enriched in stage III and IV melanosomes [5, 27, 28]. Tyrosinase is a copperdependent enzyme that catalyzes the limiting steps in melanin synthesis, ltyrosine hydroxylation to l-3,4-dihydroxyphenylalanine (l-dopa) and l-dopa oxidation to dopaquinone [29]; tyrosinase also catalyzes the downstream oxidation of 5,6-dihydroxyindole (DHI) to 5,6-indolequinone [30]. Patients with OCA type 1 have mutations in the gene encoding tyrosinase, and most often suffer from a complete loss of pigmentation in the skin and eyes [31]. As the name suggests, Dct catalyzes the tautomerization of dopachrome to DHI [32]. The murine Tyrp1 homolog has been suggested to have DHI-2-carboxylic acid (DHICA) oxidase activity [33], but the enzyme activity of the human Tyrp1 homolog has been controversial [34]; Tyrp1 has also been suggested to stabilize tyrosinase [35]. OCA type 3 results from inactivating mutations in the gene encoding TYRP1 in humans, and is characterized by accumulation of red and brown pigments [34]. Tyrosinase, Dct and Tyrp1 are type I membrane glycoproteins that share around 40% amino acid homology [36]. All three enzymes contain an N-terminal signal sequence that targets the nascent protein to the ER, a large intralumenal domain that bears the catalytic activity, a single membrane-spanning domain, and a short C-terminal cytoplasmic domain. The cytoplasmic domains of tyrosinase and Tyrp1 contain signals that are crucial for correct targeting to the melanosome [37–41]. In addition to the melanogenic enzymes, a number of other proteins required for melanogenesis are thought to be associated with melanosomes. Proteomic analyses of melanosome-enriched subcellular fractions from melanocytic cells have identiﬁed numerous proteins associated with the organelle [42, 43], but only a few of the “hits” have been veriﬁed by more rigorous analyzes. Among them are the putative channel protein, OCA2 [44], and the G protein-coupled receptor (GPCR) GPR143, commonly called OA1 [45]. These proteins are the products of genes that are targeted by mutation in patients with OCA type 2 and OA type 1, respectively. The molecular function of OCA2, also called the P protein because it is mutated in the pink-eyed dilute mouse [46], is unknown, but melanosomes in OCA2-deﬁcient melanocytes are severely hypopigmented [47, 48] and are depleted of tyrosinase [49, 50]. OCA2 is also thought to regulate melanosomal pH [51], perhaps by facilitating counteranion transport [52–54]. OA1 belongs to

9.2 Melanosomes: Intracellular Organelles Specialized in Melanin Synthesis

the large family of GPCRs with seven-transmembrane spanning domains [45, 55]. Its ligand has been recently suggested to be the melanin intermediate l-dopa [56]. OA1 has been proposed to control fusion and ﬁssion events with transport intermediates [57] as detailed in Section 9.3.4. Additional melanocyte-speciﬁc components that are likely residents of melanosomes include the potassiumdependent sodium/calcium exchanger SLC24A5 [58] and the putative transporter SLC45A2 [59]. Genetic variants of SLC24A5 are associated with skin color variation in humans [60] and mutations in SLC45A2 (also called membraneassociated transporter protein (MATP) or underwhite in mice) result in OCA type 4 [61]. Both were identiﬁed in melanosome-enriched fractions by proteomics analysis, but their predominant localization and molecular function are not yet clear. Mutations in the calcium channels TRPM1 [62] and TRPM7 [63] result in defects in melanogenesis, and thus these are also likely associated with melanosomes. Finally, the MART-1 tumor-associated antigen has been shown to interact with other melanosomal proteins, such as Pmel17 [64] and OA1 [57], and may play an important role as a chaperone in their trafﬁcking within the melanocyte. The identity and function of many of these components have been reviewed in detail elsewhere, and their deﬁciencies result in pigmentation disorders in humans (ocular or oculocutaneous albinism) and/or in model organisms (Table 9.1). Melanosomes also harbor proteins that are ubiquitously expressed and that likely play a role in melanogenesis. Most of these components, however, are not necessarily enriched only in melanosomes but may be present in other cellular compartments as well. Among these are lysosomal enzymes such as acid phosphatase and cathepsins [20, 23] and lysosomal membrane proteins such as lysosomal membrane proteins (LAMP) 1 and 2 (reviewed in [4]). Evidence from proteomics analyzes suggest that other lysosomal proteins and components of the ER might also be present in different stage melanosomes [27, 42, 43], but these ﬁndings warrant veriﬁcation by other means. One component that plays an important role in melanosomes is the copper transporter, ATP7A [65]. ATP7A is a member of the ABC family transporters that functions as an ATP-dependent pump to load copper into secretory/endocytic organelles from the cytosol [66]. Whereas in all cells ATP7A is localized predominantly to the trans-Golgi network (TGN) and endosomes, a cohort of ATP7A in melanocytes is additionally present within melanosomes, where it provides the copper cofactor required for tyrosinase activity; a failure to localize ATP7A to melanosomes results in severe hypopigmentation due to a loss of tyrosinase activity [65]. Genetic deﬁciencies in ATP7A result in Menkes disease, which is associated with severe neurological and developmental abnormalities [66]; mouse models of Menkes disease with mutations in ATP7A include the mottled and brindled mice, which were ﬁrst noticed for their effects on pigmentation [13, 67].

253

254

9 Biogenesis of Melanosomes

9.3 Endocytic System and Formation of Melanosomes 9.3.1 Organelles of the Endocytic Pathway

In most cells the endocytic pathway serves housekeeping functions such as nutrient uptake, control of signaling pathways, and macromolecule degradation [68]. These functions are carried out by exploiting distinct membrane delimited compartments and organized subdomains that are highly dynamic [69, 70]. Proteins and other macromolecules that are internalized at the plasma membrane by receptor-mediated clathrin-dependent and -independent pathways or bulk-ﬂow endocytosis [71] reach distinct compartments that are classiﬁed as “early” and “late” endosomes (Figure 9.2). This classiﬁcation is based on the sequential accessibility of these different compartments to free endocytic tracers and to ligands bound to internalized cell surface receptors. Endocytic tracers access early endosomes within 5–15 min following internalization. They consist of different domains deﬁned, morphologically, by the relative enrichment in speciﬁc effectors and by the kinetics of trafﬁcking of ligands bound

Coated pit

Early recycling endosome

Early sorting endosome

Late endosome/MVB

Lysosome

TGN Golgi Apparatus Endoplasmic Reticulum

Figure 9.2 Endocytic pathway. Schematic representation of organelles of the endosomal system, emphasizing the bestcharacterized compartments found in all cell types. Arrows represent proposed directions of protein and lipid transport between organelles. The clathrin-containing coat of the plasma membrane invagination and the

coat at the cytosolic side of the early sorting endosome (coated endosome) is represented by gray shading. Intralumenal vesicles present in early sorting endosomes and late endosomes are represented by small circles; irregularly shaped membranes and vesicles are generally present in lysosomes.

9.3 Endocytic System and Formation of Melanosomes

to internalized receptors such as those for transferrin (Tf), low-density lipoprotein (LDL), epidermal growth factor (EGF), and asialoglycoprotein (ASGP) [72]. Sorting endosomes consist of tubules that are continuous with closely apposed electronlucent vacuoles with few internal membranes, and are accessed by endocytic cargo within 5–15 min [68]. These membranes are generally associated with the small GTPase Rab5, the Rab5 effector early endosomal antigen 1 (EEA1), and the phosphorylated phosphoinositide phosphatidylinositol-3-phosphate [73]. The tubules transport cargoes into and out of the vacuoles, including receptors that rapidly recycle to the plasma membrane (e.g., Tf receptor). A proportion of such recycling receptors accumulate in and recycle from perinuclear, tubulovesicular elements that are apposed to the Golgi apparatus and are called recycling endosomes [74]. The small GTPases Rab4 and Rab11 are generally involved in the maintenance of these recycling domains, as do many other Rab GTPases (such as Rab14 and Rab22A) and their effectors (for review, see [75]), and they can accumulate recycling cargoes for up to 30 min after internalization [68]. The tubules that emanate from the vacuolar domains of early endosomes are thought to remove membranebound cargoes destined for recycling to the plasma membrane or to other secretory and endosomal compartments, while the vacuolar domains – which mature into late endosomes as these materials are removed – become enriched in soluble cargoes that are destined for degradation in lysosomes. Another sorting event that occurs from the vacuolar domains of early endosomes is the segregation of integral membrane cargoes destined for the lysosomal pathway. This sorting event is mediated by the inclusion of these cargoes on internal membrane vesicles that form by inward budding of the endosomal limiting membrane [76]. Sorting occurs from regions of the vacuolar domain limiting membrane that are adjacent to coated areas on the cytosolic face of the endosome [77]. These bilayered coats are particularly abundant in melanocytic cells [5]. This coat contains clathrin, but is distinct from clathrin coats on buds at the plasma membrane, TGN, and tubular endosomes. The coat also contains components of the so-called ESCRT (endosomal sorting complex required for transport) machinery [78]. Four ESCRT multisubunit complexes are sequentially recruited to the endosomal membrane for the sequestration of membrane proteins that are covalently modiﬁed by ubiquitin and for the formation and severing of the internal vesicles from membranes enriched in these cargoes [79]. As the vacuolar endosomes mature, they continue to accumulate internal vesicles; such maturing endosomes are called multivesicular bodies (MVBs) [76]. Fully “matured” MVBs, which typically lack associated membrane tubules, are equivalent to late endosomes. They contain LAMP1 and 2, newly synthesized lysosomal enzymes, and are accessed by ﬂuid-phase endocytic cargoes and ligand–receptor complexes within 30 min [80]. The biogenesis of MVBs and mechanisms involved in the formation of the intraluminal vesicles and sorting of cargo have been particularly well studied in yeast [76] and are largely conserved in mammalian cells during the attenuation of ligand-stimulated receptors such as the EGF receptor (EGFR) [78]. Not all MVBs are similar [81], and the mechanisms involved in the formation of distinct classes of MVBs are likely to be more complex and diverse then

255

256

9 Biogenesis of Melanosomes

originally thought. Moreover, not all MVBs are destined for fusion with lysosomes (for review, see [82]). In particular, specialized cells like melanocytes harbor an heterogeneous set of compartments with morphological hallmarks of MVBs that harbor melanosomal proteins (e.g., Pmel17) and that serve as intermediates in melanosome formation. The internal vesicles of these structures appear to form in an ESCRT-independent manner (see Section 9.3.5.3). The effectors involved in cargo sorting and the formation of these intraluminal vesicles are not yet known. While this description of the endocytic system incorporates studies from several laboratories in many cell types, one should keep in mind that endosomal membranes are highly dynamic and constantly remodeling. Distinct endosomal domains vary in prevalence or content based on cell type, differentiation state, or response to extracellular signals. As detailed below, specialized cells – including melanocytes – subcompartmentalize the ubiquitous endosomal system to facilitate regulated cargo sorting to generate cell-type-speciﬁc organelles – including melanosomes. 9.3.2 Melanosomes Are LROs but Are Distinct from Lysosomes

The unique morphology and composition of melanosomes pose a challenge for organellogenesis within melanocytes and other pigment cells. To form melanosomes, these cells must either modify a ubiquitous organelle or develop a novel mechanism for sorting speciﬁc resident proteins from ubiquitous organelles. Several observations from early immunocytochemical and subcellular fractionation studies had suggested common features for melanosomes and late endosomes/ lysosomes. These include the presence of lysosomal hydrolases and integral membrane proteins, apparent fusion with phagocytosed particles, an acidic pH (for review, see [4]) and the presence of internal vesicles like those in MVBs [83, 84]. Moreover, like lysosomal proteins in other cell types, tyrosinase activity was detected in clathrin-coated vesicles near the TGN in melanocytes [20, 85], and tyrosinase and Tyrp1 localize to late endosomes and lysosomes when expressed ectopically in transfected nonpigment cells [38, 39, 41, 86]. These features of melanosomes led to their classiﬁcation as LROs, a family of organelles that also include cytolytic granules of cytotoxic T cells, platelet α and dense granules of megakaryocytes, basophilic cell granules of mast cells, major histocompatibility complex (MHC) class II compartments in antigen-presenting cells, lamellar bodies in lung type II epithelial cells, Weibel–Palade Bodies in endothelial cells, and others [87]. Melanosomes, like other LROs, were once considered as transformed lysosomes that act as dual-functional organelles, carrying out both specialized and universal functions [4]. However, it is now clear that all LROs do not necessarily share common biogenetic pathways. While some LROs (such as perhaps cytolytic granules [88]) appear to be modiﬁed lysosomes, others (including melanosomes, platelet dense granules, and lamellar bodies) coexist with bona ﬁde late endosomes/

9.3 Endocytic System and Formation of Melanosomes

lysosomes and thus must derive separately from other endocytic and biosynthetic organelles. This distinction explains why LROs are not uniformly disrupted in genetic disorders such as HPS (see Section 9.4) [89]. Evidence that melanosomes coexist with conventional lysosomes – and thus likely represent a separate organelle lineage – was supported by early histochemical observations. Seiji et al. reported that while lysosomal acid phosphatase activity was detected histochemically in melanosomes of melanoma cells, nonmelanosomal structures similar to lysosomes contained higher activity [21]. Moreover, studies from Boissy et al. showed that acid phosphatase activity was absent from melanosomes and premelanosomes of untransformed skin melanocytes [90]. These early observations were extended by studies from our group in skin melanocytic cell models using quantitative immunoelectron microscopy [5]. These analyzes showed that only about 10–20% of cellular LAMP1 and cathepsin D was present in melanosomes, whereas the vast majority of these proteins was detected in separate structures with morphological and temporal characteristics of lysosomes. These compartments were more acidic than mature melanosomes based on the accumulation of the weak base (3-(2,4-dinitroanilino)-3′-amino-Nmethyldipropylamine) (DAMP) generally used as a probe to determine the acidity of intracellular compartments [91]. Moreover, using endocytic tracers such as goldconjugated bovine serum albumin, we showed that stage I melanosomes were accessible to the endocytic system shortly after endocytosis, but that stage II–IV melanosomes were not. These observations indicated that melanosomes diverge and segregate from the endosomal system at the level of stage I melanosomes. These ﬁndings have been veriﬁed by others [92] and supported by numerous studies in melanocytes from disease models such as HPS (see Section 9.4.2). By contrast, one study in cultured choroidal melanocytes suggested that melanosomes are accessible to endocytic tracers [93], but this might reﬂect extended incubations of cells with tracers that induce autophagic processes. In other studies, tyrosinase or Tyrp1 were detected largely in subcellular fractions with lysosomal enzyme activity [23, 94, 95], but the “melanosomal” fractions were not assessed for purity and may have contained late endosomes and/or lysosomes. All together the evidence supports the conclusion that eumelanosomes are not merely modiﬁed lysosomes, and that sorting mechanisms within pigment cells distinguish between lysosomal and melanosomal cargoes (Figure 9.3). In addition to the differences in protein contents, melanosomes and lysosomes differ in pH. Using DAMP accumulation as an indicator for low pH, premelanosomes are highly acidic, accumulating nearly as much DAMP as lysosomes, but progression from premelanosomes to mature melanosomes is accompanied by a decrease in acidity [5]. This alkalization may be driven by the progressive removal of a vacuolar ATPase during melanosome maturation or by inactivation of a premelanosome-speciﬁc proton pump, such as the OCA2 transporter [51]. Tyrosinase is inactive below pH 5 [96, 97]. Thus, a progressive increase in pH from early to late melanosomes might prevent premature tyrosinase activity within earlystage melanosomes but favor higher activity, and consequently facilitate melanin biosynthesis, as melanosomes mature.

257

258

9 Biogenesis of Melanosomes

Tyrosinase Tyrp1 Tyrp2/DCT

Early recycling endosome

Pmel17

Stage II

Stage III

Stage IV

Stage I

Early sorting endosome TGN Golgi Apparatus

Late endosome/MVB

Figure 9.3 Melanosomes originate within the endosomal system, but are distinct from lysosomes. Schematic representation of the endocytic and melanosomal organelles, and a working model for the sorting pathways for proteins distributed among them. Pmel17 is delivered to early sorting endosomes either following endocytosis from the plasma membrane or directly from the TGN. Domains of these endosomes then mature into coated endosomes/stage I melanosomes with characteristic internal vesicles and short ﬁbrils and then into stage II melanosomes in which the ﬁbrils organize

Lysosome

into sheets. By contrast, Tyrp1 is delivered to maturing melanosomes from an endosomal intermediate to which it is targeted directly from the TGN. Tyrosinase is also targeted to melanosomes from an endosomal intermediate. Delivery of the melanosomal enzymes results in maturation of the stage II premelanosome to the stage III and IV melanosome. The deposition of black melanin in these compartments is schematized by bold lines on the top of the striations (stage III) or by a black content throughout the organelle (stage IV).

It is likely that the segregation of melanosomal and lysosomal proteins is regulated by melanocyte differentiation and/or by differences in cell type, tissue of origin, or degree or type of pigmentation. In support of this notion, melanosomal proteins like tyrosinase, Tyrp1, OCA2, OA1, and Pmel17 localize to late endosomes/ lysosomes when expressed in nonpigmented cells [38, 39, 41, 44, 86, 98, 99]. Furthermore, different cohorts of melanosomal proteins are expressed in melanocytes during synthesis of pheomelanins and eumelanins [100, 101], and melanosomal proteins are less efﬁciently segregated from lysosomal proteins in hypopigmented melanocytic cells or brown melanocytes (G. Raposo and M.S. Marks unpublished data). RPE cells have morphologically distinct melanosomes (for review, see [102]) that may fuse more readily with lysosomes. A developmental difference in lysosome/melanosome segregation could explain why phagocytosed latex particles could be observed in melanosomes of RPE cells and choroidal melanocytes [103],

9.3 Endocytic System and Formation of Melanosomes

whereas endocytic tracers failed to access premelanosomes or melanosomes in skin-derived melanocytic cells. 9.3.3 Pmel17 and Generation of Early-Stage Melanosomes

Stage I and II melanosomes lack pigment, and thus are enriched in a distinct set of proteins from those that function in melanogenesis and that are enriched in stage III and IV melanosomes. The most prominent characteristics of early-stage melanosomes are the intralumenal ﬁbrils that mature into parallel sheet-like arrays. These ﬁbrils appear to consist primarily of fragments derived from a single protein, Pmel17. The maturation of Pmel17 from an integral membrane protein to a ﬁbrillar form has been singularly instructive in understanding how early-stage melanosomes form. As discussed below, it has also provided surprising and novel insights into functional amyloid formation. A brief discussion of Pmel17 maturation is therefore warranted here. 9.3.3.1 Pmel17 Structure Like the melanin biosynthetic enzymes tyrosinase, Tyrp1, and Dct, Pmel17 is synthesized as a type I transmembrane protein with a short N-terminal signal sequence, a large lumenal domain, a single membrane spanning domain, and a short cytosolic domain [25]. In humans, two independent alternative mRNA splicing events generate four different Pmel17 protein products that differ within the lumenal domain [104–107], functional differences for these isoforms have not yet been deﬁned. The lumenal domain can be subdivided into four distinct subdomains based on sequence homologies: an N-terminal region (NTR) with no known structural homolog, a PKD domain with homology to a repeated element within the polycystic kidney disease-1 protein, a RPT (proline/serine/threoninerich repeat) domain consisting of 10 imperfect direct repeats of a 13-residue sequence, and a membrane-proximal cysteine-rich Kringle-like domain (KLD) (reviewed in [25]). Hearing et al. further subdivide the lumenal domain to include gap regions (GAP1, GAP2, and GAP3) between the NTR and PKD, between the RPT and KLD, and between the KLD and transmembrane domains [108, 109]; here we will use the more simpliﬁed terminology. Pmel17 expression is normally limited to pigment cells. In humans Pmel17 expression is maintained in most melanoma cells [110, 111], but can also be induced in unusual “clear cell” tumors of mixed lineage [112, 113]. Hence, human Pmel17 has been well studied as a tumor-associated antigen, and numerous reagents – including antibodies and T cell clones – have been developed to detect it. These antibodies have proven to be extraordinarily useful in detailing the intracellular itinerary of Pmel17 as it matures within the melanocyte and in detecting Pmel17 by immunoelectron microscopy. 9.3.3.2 Pmel17 Forms the Fibrillar Matrix upon Which Melanins Deposit Unlike other melanosomal proteins discussed earlier, Pmel17 is most easily detected – using monoclonal antibodies to the lumenal domain – in stage I and II

259

260

9 Biogenesis of Melanosomes

melanosomes and becomes less easily detected in stages III–IV [5, 114]. Orlow’s group ﬁrst suggested that Pmel17 might be part of the “melanosome matrix” (i.e., the intralumenal ﬁbrils) based on its insolubility in aqueous nonionic detergents or in the detergent phase of Triton X-114 [115, 116]. Consistently, immunolabeling for Pmel17 within stage II melanosomes was associated with the ﬁbrils [5] and puriﬁed ﬁbril-enriched subcellular fractions from melanocytic cells contain Pmel17 fragments [117]. Pmel17 immunoreactivity by microscopy decreases as melanosomes mature to stage III and IV [5], and biochemical detection of Pmel17 is inhibited by melaninization [118], consistent with masking of the Pmel17 ﬁbrils by polymerized melanins during melanosome maturation. Pmel17 is the only pigment cell-speciﬁc protein required to generate the ﬁbrils in vertebrate cells, as shown by the arrays of melanosome-like ﬁbrils detected by electron microscopy in endosomes of transfected nonmelanocytic HeLa cells ectopically expressing Pmel17 [98]. Pmel17 is also necessary for ﬁbril formation in melanocytes, as shown by analyses of melanocytic cell lines derived from silver mice in which the Pmel17 cytoplasmic domain is truncated [119]. Finally, recombinant fragments generated in bacterial expression systems and derived from the Pmel17 lumenal domain form premelanosome-like ﬁbrils in vitro [120, 121]. Together, these studies suggest that Pmel17 is the main – if not sole – component of the premelanosome ﬁbrils. Importantly, the ﬁbrils bear biophysical hallmarks of amyloid [120, 121] – a repeating β-sheet-rich protein fold normally associated with misfolded proteins in neurodegenerative diseases such as Alzheimer’s and Parkinson’s disease [122]. Pmel17 thus represents the ﬁrst of a class of mammalian “functional amyloids” and a model for the formation of pathological amyloids [123]. 9.3.3.3 Pmel17 Biosynthesis and Amyloid Formation How does a transmembrane protein like Pmel17 form amyloid-like ﬁbrils solely within the lumen of stage I–II melanosomes? A key to understanding this remarkable protein transformation is in the intracellular journey of Pmel17 from its point of synthesis in the ER to stage I melanosomes. The results of a number of studies (reviewed in [25] and [124]) paint the following picture of Pmel17 maturation. Pmel17 is synthesized in the ER as a precursor (P1) form with four core N-linked glycans. Three of the N-linked glycans become processed to the mature form upon passage through the Golgi complex. In addition, Pmel17 is extensively modiﬁed by O-linked glycosylation within the RPT domain and these oligosaccharides are further modiﬁed in the Golgi, including terminal sialic acid (at least in cell culture). Sialylated O-linked oligosaccharides are critical for Pmel17 recognition by the monoclonal antibody HMB45 [125–129], which is often used as a reagent to detect melanoma [126]. The full-length Golgi-matured form of Pmel17 has been referred to as the precursor 2 (P2) form [98]. From the Golgi, at least a fraction of Pmel17 traverses the plasma membrane, where it is internalized into the endosomal system by clathrin/AP-2-dependent endocytosis interacting with a di-leucine-based endocytosis signal within the Pmel17 cytoplasmic domain [119, 130, 131]; a natural truncation mutant of Pmel17 in the silver mouse removes this signal, resulting in

9.3 Endocytic System and Formation of Melanosomes Stage I

Amyloid fibrils Pmel17

Stage II

Mβ Mα

Model for the formation of premelanosome ﬁbrils. The melanocytespeciﬁc protein, Pmel17, is present on the limiting membrane and the internal vesicles of early coated endosomes (stage I melanosomes). Within these structures, Pmel17 is cleaved to Mα (purple/orange lumenal domain) and Mβ fragments (blue cytoplasmic domain) by a proprotein convertase (represented by a scissor). Mα fragments that dissociate from membranes are subsequently cleaved by still unknown proteases and begin to assemble in small, irregular ﬁbrils that become fully organized

Figure 9.4

within stage II melanosomes. Only Mα and derived fragments (top right) are reproducibly detected in stage II melanosomes. On the bottom right, a three-dimensional model of a stage II melanosomes obtained by tomographic reconstruction of a region of the cytoplasm of an MNT-1 melanoma cell preserved by high-pressure freezing before freeze substitution and embedding in Epon. Stage II premelanosomes (membrane in pink) bear amyloid ﬁbrillar sheets (yellow/ gold) and intraluminal vesicles (green). Scale bars: 200 nm.

accumulation at the plasma membrane and depletion from melanocytes [119]. Within endosomes or perhaps the TGN, Pmel17 is cleaved at two speciﬁc sites within the lumenal domain by two proteases (Figure 9.4). First, a proprotein convertase cleaves Pmel17 at residues 468–469 (human Pmel17), separating Pmel17 into two disulﬁde-linked fragments: a large lumenal fragment referred to as Mα (mature polypeptide α) – containing the NTR, PKD, and RPT domains – and a smaller, membrane bound fragment called Mβ (mature polypeptide β) – containing the KLD, transmembrane, and cytoplasmic domains [98, 117]. Second, a “site 2 protease” cleaves Mβ within the juxtamembrane region of the lumenal domain to liberate Mα and its associated lumenal part of Mβ from the membrane [132]; the identity of this protease is not yet clear, but depletion of either of two proteases of the ADAM (“a disintegrin and metalloprotease domain”) family of α-secretases

261

262

9 Biogenesis of Melanosomes

severely impair this cleavage and downstream events [132]. Indirect evidence suggests that the cleavages occur within speciﬁc domains of early endosomes [119], and both seem to require the acidic environment within the late secretory and endocytic pathways [98] (and B. Watt and M.S. Marks, unpublished results). A small fraction of the cleaved forms of Pmel17 is secreted from cells in culture [133, 134], but the functional signiﬁcance of this cohort is not yet known. The membrane-bound fragment of Mβ remaining after site 2 protease cleavage is a substrate for intramembrane proteolysis by the γ-secretase complex [132], well known for its activity on the Alzheimer precursor protein, Notch, and other clinically important substrates [135]. Once cleaved and released from the membrane within endosomes, Pmel17-Mα has the unusual property of aggregating in an ordered fashion into polymerized ﬁbrils. Indeed, it is Mα (or fragments derived therefrom) that polymerizes into amyloid in vitro [120, 121, 136] and Mα released that is released from Mβ is only detected within fractions of melanocytic cells that are insoluble in nonionic detergents [117]. Fibrillogenesis requires all three subdomains of Mα [108, 119]. However, Mα is only a minor species detected within these fractions. Rather, Mα is further digested to smaller fragments corresponding at least to the RPT and part of the PKD domains [27, 108, 121, 129]. These are the fragments that are detected by immunocytochemistry on stage II melanosomes, and that are likely buried by melanins in stages III and IV; epitopes corresponding to the N- and C-termini of Pmel17 are not detected in melanosomes, likely due to proteolysis (indeed, antibodies to the C-terminus follow Mβ and its derived C-terminal fragment and are excluded from stage II–IV melanosomes) [5, 98, 129]. Thus, the progression of Pmel17 through the secretory pathway and its regulated cleavage within acidic endosomal compartments accounts for the ability of this endogenously synthesized protein to assemble into ﬁbrils only at a given stage of the endosomal pathway. 9.3.3.4 Functional Importance of Fibrillar Melanosomes What is the function of the Pmel17 amyloid ﬁbrils? The answer is not entirely clear. Pmel17 ﬁbrils – and indeed, any amyloid ﬁbril – accelerates melanin polymerization in vitro [106, 114, 120], suggesting that Pmel17 might play a kinetic role in melanization or in absorbing otherwise toxic oxidative melanin intermediates. Pmel17 is responsible for the elliptical shape of melanosomes, as silver mice have round, enlarged melanosomes [119]; this shape might be important for melanosome transfer from melanocytes to keratinocytes or for melanosome motility into the apical processes of RPE [137]. Lastly, the polymerization of melanins into a solid mass composed of ﬁbrillar sheets rather than smaller, separate packages might facilitate melanin transfer to keratinocytes if melanins are released from melanocytes by secretion. However, it should be noted that melanocytes from silver mice are highly pigmented and that the coatcolor defect in these mice is mild [13, 67, 138]. Elucidation of the true function of these ﬁbrils will need to await a complete Pmel17 gene deletion in a tractable experimental system.

9.3 Endocytic System and Formation of Melanosomes

9.3.4 OA Type 1 and Melanosome Biogenesis

The OA1 protein is a pigment cell-speciﬁc member of a subfamily of GPCRs [55, 139, 140]. Mutations in the OA1 gene underlie OA type 1, an X-linked disorder that does not impair melanin production but decreases the number of melanosomes in RPE and the choroid [141]. Although skin is pigmented in OA1 patients and mouse models, aberrant giant melanosomes, called “macromelanosomes,” accumulate in both RPE and skin melanocytes, suggesting a defect in melanosome biogenesis [142]. Like other canonical GPCRs, OA1 interacts with arrestins and activates heterotrimeric G-proteins [45, 55, 140]. Null mice for the inhibitory G-protein Gαi3 show similar phenotypes to those observed on OA type I albinism, suggesting that OA1 might control melanosome maturation by activating the G-protein Gαi3 [143]. Unlike other GPCRs that localize mainly to the cell surface, OA1 localizes intracellularly to lysosomes and melanosomes [57, 99] by virtue of speciﬁc sorting signals in its cytosolic domain [99]. Thus, whereas other GPCRs bind extracellular ligands, the OA1 ligand is likely exposed in the lumen of the melanosome or other compartments. One report has suggested that l-dopa – the melanin precursor – is the OA1 ligand [56]. Ligand binding triggers a signaling cascade from the organelle lumen to the cytosol. While this cascade remains poorly characterized, the phenotype of OA1 mutant cells predicts that it signals organelle-intrinsic fusion/ﬁssion events during melanosome biogenesis, similar to the phagosome intrinsic regulation of antigen processing by Toll-like receptor signaling in dendritic cells [144]. The membrane trafﬁcking steps regulated by OA1 have remained enigmatic. Recent studies by our group suggest that the giant melanosomes observed in OA1deﬁcient melanocytic cells result from aberrant fusion/ﬁssion events at early steps of melanosome biogenesis [57]. Transient OA1 downregulation in a eumelanogenic melanoma cell line results in the generation of aberrant premelanosomes that accumulate cargo proteins of immature melanosomes (Pmel17), late-stage melanosomes (Tyrp1), and lysosomes (LAMP1), suggesting that protein sorting and/or segregation of organelles downstream of coated endosomes is somehow regulated by OA1 [57]. Another recent study suggested that OA1 might function not only in the regulation of melanosome biogenesis, but also in melanosome motility. This study showed that melanosomes in skin melanocytes and RPE from Oa1-null mice are abnormally distributed toward the cell periphery [145]. It is possible that both phenotypes are linked, such that the altered motility is a consequence of improper melanosome maturation (for review, see [146]). 9.3.5 Origin of the Melanosome 9.3.5.1 Early-Stage Melanosomes Originate within the Endocytic Pathway Having deﬁned Pmel17 as a major biogenetic constituent of early-stage melanosomes provided a means with which to probe the origins of these unusual organelles within the secretory/endosomal system. Overwhelming evidence now

263

264

9 Biogenesis of Melanosomes

supports an early endosomal origin for stage I and II melanosomes. While the details of how these organelles diverge from endosomes is not yet complete, our observations thus far have provided a new paradigm for LRO biogenesis, and highlighted the endocytic system as a unique and highly adaptable platform from which multiple sorting pathways can converge to generate the melanosome. Compelling evidence for an early endosomal origin of the premelanosome came ﬁrst from immunoelectron microscopy analyzes of Pmel17 distribution in eumelanogenic cells. A cohort of mature Pmel17 was detected in EEA1-positive early endosomal tubules and vacuoles [5]. Some of the vacuoles displayed bilayered clathrin coats on the cytosolic side. These compartments clearly correspond to vacuolar sorting endosome domains based on the presence of EEA1, the ﬂat bilayered coats, a few intralumenal membranes, and accessibility to internalized Tf and gold-conjugated bovine serum albumin after short incubations [5]. These compartments are also structurally equivalent to the stage I premelanosome [5, 10] and are early endosome precursors to stage II premelanosomes based on several lines of evidence, including: (i) the progressive loss of EEA1 correlates with a progressive increase of Pmel17 density during progression from tubular early endosomes to coated endosomes to stage II premelanosomes; (ii) endocytic tracers progressively access early endosomes and then coated endosomes, and are excluded from stage II premelanosomes; and (iii) early and coated endosomes, but not the stage II premelanosome, are labeled by an antibody to the C-terminus of Pmel17, which becomes segregated from the ﬁbrillinogenic lumenal domain after cleavage [5, 117]. Together, these observations suggest that Pmel17 passes through the early endosome and the coated endosome intact before its deposition as ﬁbrils in the stage II premelanosome. Within coated endosomes, Pmel17 accumulates largely on intraluminal membranes, similar to integral membrane cargoes destined for lysosomal degradation [5]. In fact, when expressed in nonpigment cells, Pmel17 accumulates primarily on intraluminal membranes of MVBs [98]. Several lines of evidence support the notion that Pmel17 begins to form ﬁbrils in association with these intraluminal vesicles. First, in transfected nonpigment cells that express Pmel17, ﬁbrils form within MVBs intermingled with the intraluminal membranes [98, 117]. Second, inhibition of sorting onto the intraluminal ﬁbrils by mutagenesis of the Pmel17 NTR or PKD domains ablates ﬁbril formation – as well as the proprotein convertase cleavage required for liberating the ﬁbrillogenic Pmel17-Mα fragment [147]. Third, and most directly, ﬁbrils are seen to emanate from intralumenal vesicles in threedimensional reconstructions from electron tomographic analyzes of coated endosomes/stage I melanosomes in a eumelanogenic melanoma line [11]. Using cryopreservation rather than chemical ﬁxation, intermediates could be visualized ranging between coated vacuolar endosomes with small protoﬁbrils and elongated stage II melanosomes with ﬁbrillar sheets [11]. These observations conﬁrm a direct role for the intraluminal membranes as “seeds” for the formation of Pmel17 amyloid during the biogenesis of early-stage melanosomes. Intraluminal membranes of multivesicular endosomes have also been implicated in the formation of pathogenic amyloid by the Alzheimer’s Aβ peptide and the prion protein [148],

9.3 Endocytic System and Formation of Melanosomes

suggesting that Pmel17 and pathogenic amyloidogenic proteins use a common mechanism in their transformation into ﬁbrils. 9.3.5.2 Melanosomes Do Not Originate from the ER The notion that early-stage melanosomes develop from endosomes has been challenged by an alternative view that stage II melanosomes emerge from the ER. However, the precedents on which this view is based are ﬂawed. The idea originated from an elegant early study by Maul, who used electron microscopy analyses of serial sections of a human melanoma line to show that structures with the characteristic morphologic features of stage II melanosomes could be observed in continuity with smooth tubular membranes near the Golgi apparatus [149]. Maul used the term “smooth ER” to describe these membranes, because at the time this was the only “smooth” tubular membrane system known; the TGN and the endocytic system had not yet been characterized morphologically. The membranes were not further identiﬁed; in fact, it is likely that they corresponded to endosomal tubules, which we now understand can fuse with immature melanosomes during maturation (see Section 9.3.5.4). Nevertheless, Hearing et al. have proposed an origin for stage I/II melanosomes from the rough ER based largely on subcellular fractionation of melanocytic cells and immunoblotting analyses. A subcellular fraction of a human eumelanogenic melanoma line that was enriched in stage II melanosomes was shown to contain the full-length, unprocessed P1 form of Pmel17 by immunoblotting using antibodies to the cytoplasmic domain. Maturation from stage II to stage III melanosomes was accompanied by a loss of this form and an accumulation of RPT domaincontaining proteolytic fragments detected by monoclonal antibody HMB45 [27]. Moreover, proteomic analysis of this fraction revealed the presence of traditional ER resident proteins in addition to known constituents of early and late-stage melanosomes, late endosomes/lysosomes, lipid rafts, and secretory granules [42]. While the authors inferred that stage II melanosomes contain ER proteins and stem from the ER, these observations are more likely explained by contaminating ER membranes within these fractions (for a discussion, see [25]). The inability to detect Golgi-processed forms of Pmel17 by immunoblotting with the antibody to the C-terminus in these and other studies [27, 150, 151] was taken as further evidence for a direct formation of melanosomes from the ER. However, these analyses failed to take into account the rapid processing of Pmel17 compared with the slow folding and release from the ER [98], the loss of the C-terminus during maturation [98, 117], and the detergent insolubility of ﬁbrils [108, 117, 129, 152, 153], which together make the P1 form the most predominant by far at steady state within detergent-soluble cell fractions. Moreover, as discussed earlier, HMB45 reactivity with Pmel17-derived bands detected by these authors in stage III melanosome-enriched fractions requires processing by Golgi enzymes. Finally, signal overlap by immunoﬂuorescence microscopy has been used as an argument that the Pmel17 C-terminus and HMB45-reactive ﬁbrils are detected in the same compartment [152, 153], but these analyses failed to account for the wide distribution of the ER throughout the cell or to provide higher resolution views – as in

265

266

9 Biogenesis of Melanosomes

other studies [5, 129] – in which the ER and stage II melanosomes could be resolved. Thus, the evidence overwhelmingly supports an origin within the endocytic system. Nevertheless, a role for the ER during the melanosome maturation process should not be disregarded. The generation of melanosomes requires functions that could potentially be effected through speciﬁc interactions between maturing melanosomes and the ER, including lipid metabolism, calcium homeostasis, and redox reactions. Tight interactions between the ER and melanosomes could even potentially explain the presence of ER components in isolated fractions of melanosomes [42, 43], as has been shown for phagosomes [154]. 9.3.5.3 Melanosomes Segregate from the Endocytic Pathway beyond Stage I Melanosomes Whereas endocytic tracers such as bovine serum albumin access coated endosomes/ stage I melanosomes, they do not appear to access mature melanosomes (stages II–IV), but rather pass through late endosomes en route to lysosomes with kinetics similar to those in other cells [5]. The presence within the coated endosome of both endocytic tracers and Pmel17 at early timepoints and their segregation at later timepoints indicates that this compartment is a critical sorting point between the endocytic and premelanosomal pathways. This sorting may be required for the segregation of proteins required for the morphogenesis and function of specialized endosomal organelles like the melanosome. The segregation from endosomes, rather than from the TGN, may reﬂect requirements for (i) the formation of MVBs in premelanosome biogenesis (see Section 9.3.1) and (ii) acidiﬁcation through a vacuolar ATPase in organelles such as premelanosomes. The latter is supported by the efﬁcient accumulation of the weak base DAMP in coated endosomes and stage II premelanosomes [5] and the disruption of melanosome biogenesis by inhibitors of the vacuolar ATPase ([98] and our unpublished observations). Although coated endosomes appear morphologically similar to vacuolar endosomes in which ubiquitylated cargoes are sorted to intraluminal vesicles, the sorting mechanism appears to be different. In melanocytic cells in which the ESCRT pathway was blocked by a variety of means, a traditional ubiquitylated substrate – the melanocyte-speciﬁc protein, MART-1 [155] – was protected from degradation and excluded from internal vesicles, but Pmel17 was still sorted to internal vesicles [147, 156]. Moreover, whereas mutagenesis of all ubiquitylation acceptor sites on the Pmel17 cytoplasmic domain had no effect on its sorting to internal membranes, sorting was completely inhibited by deletion of either of two lumenal subdomains [147]. The mechanism by which Pmel17 becomes incorporated onto the internal membranes remains to be determined, but might be related to its property to aggregate or to potential interactions with lipids such as ceramide [157]. The segregation of Pmel17 and MART-1 into different intralumenal vesicles suggests separate sorting for cargoes fated to degradation or to the melanosomal lineage at the level of the coated endosome. It is tempting to propose that the clathrin-containing coats act as platforms to facilitate the clustering of proteins bound for late endosomes and lysosomes, permitting concentration of Pmel17 to

9.3 Endocytic System and Formation of Melanosomes

levels necessary for ordered aggregation. A similar mechanism has been proposed for the concentration of insulin and other cargoes in nascent secretory granules [158] and in maturing transcytotic vesicles [159]. 9.3.5.4 Components of Mature Melanosomes Are Sorted from Distinct Endosomal Intermediates Melanosomal enzymes and transporters such as tyrosinase, Tyrp1, OCA2, and ATP7A are not detected in stage I and stage II premelanosomes, but are enriched in stage III and stage IV melanosomes [5, 44, 65]. This suggested that cargo sorting to mature melanosomes is a distinct process from cargo sorting to early-stage melanosomes and likely occurs from different sites in pigment cells. The cytochemical detection of tyrosinase activity in clathrin-coated vesicles near the TGN [85], the detection of tyrosinase, Tyrp1 and Dct in clathrin-coated vesicle-enriched subcellular fractions [33], and the colocalization of Tyrp1 and the lysosomal protein LAMP1 in vesicular structures near the TGN [5] led to the proposal that melanosomal proteins might be directly targeted to melanosomes from the TGN [5]. However, more recent results provide strong evidence that these cargoes are trafﬁcked to maturing melanosomes via post-Golgi endosomal intermediates. These studies have exploited either cultured melanocytes derived from mouse models of trafﬁcking disorders such as HPS or genetic manipulation of cultured melanocytes or melanoma cells to interfere with endosomal transport. As discussed further in Section 9.4.2, the products of the genes that are defective in HPS localize primarily to endosomal compartments and mature melanosomal cargoes in these models become trapped in endosomal intermediates, whereas stage II melanosomes are largely unimpaired. Evidence from analyses of wild-type and genetically altered melanocytic cells (see Section 9.4) suggests that cargoes are delivered to melanosomes from endosomes via two distinct classes of intermediates – vesicles that bud from tubular endosomal domains, and tubules that make direct contacts between endosomal vacuoles and melanosomes (Figure 9.5). Whereas vesicles have been thought to be the main mediators of anterograde trafﬁc within the endosomal system [160], direct tubular contacts between endosomal organelles have been observed in some cases [161] (Figure 9.5). In melanocytes, direct tubular contacts between vacuolar endosomes and maturing stage III/IV melanosomes have been observed using both three-dimensional reconstructions from electron tomography analyses and live cell imaging [162]. These tubules contain cargoes of recycling endosomes and thus likely correspond to specialized recycling endosomal domains that derive from the vacuoles. Membrane continuities between the tubules and both organelles suggest that they transiently fuse with melanosomes, and thus likely allow for the exchange of contents between endosomes and melanosomes [162]. Consistently, accumulating evidence suggests that at least one major cargo transport pathway from endosomes to maturing melanosomes requires components known to regulate recycling endosomal sorting and positioning (see Section 9.4.2.1). Therefore, the data suggest that these tubules are specialized endosomal domains that serve as conduits for the delivery of cargoes to maturing stage III/IV melanosomes.

267

268

9 Biogenesis of Melanosomes Melanosome

Melanosome

Recycling endosome

Figure 9.5 Endosomal tubules and buds are close to and continuous to melanosomes. (A) Electron micrograph of an ultrathin cryosection of a MNT-1 cell showing tubular membranes harboring coated buds (small arrows) closely apposed to a melanin containing melanosomes. (B) Schematic representation of the endosomal– melanosomal network, with clathrin coats represented by gray shading. (C) Threedimensional model obtained after tomographic reconstruction of a region of the

cytoplasm of a MNT-1 cell ﬁxed by highpressure freezing before freeze substitution and embedding in Epon. Left panel: tubular membranes (green) that are continuous with the melanosome limiting membrane (red) are indicated by arrows. In white, the ER. On the right panel are shown two different orientations of the three-dimensional model highlighting continuities between the endosome and the melanosome (arrows). Scale bars: 200 nm.

As melanosomes mature from stage II to stage IV, melanosome size is maintained relatively constant. Thus, the delivery of membrane to melanosomes by anterograde fusion events that facilitate cargo delivery must be balanced by retrograde transport out of melanosomes. It is possible that the endosomal tubules observed between endosomal vacuoles and melanosomes provide for both anterograde and retrograde events. Indeed, these carriers harbor endosomal cargoes that do not accumulate in melanosomes [162]; thus, either the entry into melanosomes is “gated” such that only certain cargoes are permitted to ﬂow from the tubule membrane into melanosomes or the bulk ﬂow of membrane through these tubules is from the melanosome to the endosome, with only specialized cargoes transferring in the other direction. Future live cell imaging analyses will be required to differentiate between these views.

9.4 Melanosome Maturation: Cargo Sorting to Mature Melanosomes

9.4 Melanosome Maturation: Cargo Sorting to Mature Melanosomes

Most known components of mature melanosomes are integral membrane proteins that bear cytoplasmic targeting signals required for their delivery to melanosomes [37–39, 41, 44, 99]. What cellular components recognize these signals and coordinate the intracellular transport events required to accumulate these proteins speciﬁcally within melanosomes? Key clues have come from analyses of cells derived from patients and mouse models in which OCA is one component of a syndrome of disorders characterized by generalized defects in LROs. Other clues derive from inhibitory RNA approaches in model cultured melanocytic cells (Table 9.2). 9.4.1 Griscelli Syndrome and CHS

Among diverse pathological conditions that disrupt lysosomes and related organelles there are three genetic disorders that affect melanosomes: Griscelli syndrome (GS), CHS, and HPS. GS and corresponding mouse models have been invaluable to decipher the molecular mechanisms that underlie the motility of melanosomes and other LROs [163]. Genetic, cellular, and biochemical analyses of GS-derived melanocytes revealed that an actin-based motor protein, myosin Va, is recruited to mature melanosomes by the concerted action of a member of the Rab family of small GTPases, Rab27a, and the Rab27a effector, melanophilin. These three proteins coordinate melanosome translocation from microtubules to actin ﬁlaments in the cell periphery for their ultimate transfer to keratinocytes [163, 164]. A similar complex with a different motor (myosin VIIa) and Rab27a effector (Myrip) regulate diurnal light cycle-dependent melanosome motility in mouse retinal pigment epithelia [137] (for further details, see Chapter 10). CHS is a rare genetic disorder of multiple organ systems characterized minimally by severe immunodeﬁciency, neuropathy, and OCA. The gene targeted in CHS (and the corresponding mouse beige mutation in mouse) encodes an approximately 400-kDa protein cytosolic protein called LYST (lysosomal trafﬁcking regulator) or CHS1p [165, 166]. Many cell types from CHS patients or beige mice harbor giant lysosomes and LROs (including melanosomes in melanocytes), indicating a defect in organelle biogenesis, but the exact function of LYST within pathways of organelle formation is not yet clear. Owing to its unusually large size, LYST is difﬁcult to express in physiologically relevant models. Dominant-negative approaches suggest that LYST might play a role in phosphoinositide metabolism [167]. Consistently, our studies in B lymphocyte cell lines derived from CHS patients suggest that LYST is required for sorting endosomal resident proteins into multivesicular endosomes [168]. Since multivesicular endosomes are also intermediates in melanosome formation, defects in sorting to MVBs are likely to impact melanosome biogenesis. More work needs to be done to deﬁne how LYST might regulate this process in pigment cells and whether CHS reﬂects a defect in the formation of early-stage melanosomes, late stage melanosomes, or both.

269

270

9 Biogenesis of Melanosomes

Table 9.2 Molecular complexes and proteins playing key roles in melanocyte trafﬁcking.

Intracellular trafﬁcking protein

Organelle localization

Function

Human disease/mouse model

BLOC-1

early endosome (tubules)

trafﬁcking from endosomes to melanosomes

HPS7 (h)–HPS8 (h)/muted (m)–pallid (m)–reduced pigmentation (m)–capuccino (m)–sandy (m)

Tyrp1–ATP7A BLOC-2

BLOC-3

AP-3A

RABGGTaseII

early endosome (tubules)

trafﬁcking from endosomes to melanosomes

melanosome

vesicular trafﬁcking

Golgi vesicles

Tyrp1–tyrosinase

early endosome (buds)

trafﬁcking from endosomes to melanosomes

cytosol

prenylation of Rab

Tyrp1–tyrosinase?

HPS3 (h)–HPS5 (h)–HPS6 (h)/cocoa (m)–ruby eye2 (m)–ruby eye (m) HPS1 (h)–HPS4 (h)/pale ear (m)–light ear (m) HPS2 (h)/pearl (m)–mocha (m)

interacts with cytoplasmic tail of tyrosinase HPS-like phenotype gunmetal (m)

HOPS (VPS33A)

cytosol

fusion events

HPS-like phenotype

tethering factor for SNAREs

Rab38

Rab32

Tyrp1

buff (m)

melanosome

trafﬁcking from TGN/endosomes to melanosomes

HPS-like phenotype in rats/chocolate (m)

post-Golgi tubular membranes

Tyrp1–Tyrosinase

unknown

trafﬁcking from TGN/endosomes to melanosomes

unknown

Tyrp1–tyrosinase AP-1A

KIF13A

RAB7

early endosome (buds)

trafﬁcking from endosomes to melanosomes

early endosome (tubules)

trafﬁcking from endosomes to melanosomes

unknown/embryonic lethal in mice

interacts with cytoplasmic tail of Tyrp1–tyrosinase unknown

interacts with AP-1 and Tyrp1

melanosome

vesicular trafﬁcking

late endosome

Tyrp1

unknown

9.4 Melanosome Maturation: Cargo Sorting to Mature Melanosomes

271

Table 9.2 (Continued)

Intracellular trafﬁcking protein

Organelle localization

Function

Human disease/mouse model

TSG101 (ESCRT-I)

endosomes

component of the ESCRT machinery

unknown

trafﬁcking from endosomes to melanosomes Tyrp1 Myosin Ib

endosomes

actin-associated motor

unknown

facilitates endosomal sorting of Pmel17 BLOC: biogenesis of lysosome-related organelle complex; HPS: Hermansky – Pudlak syndrome; RABGGTaseII: Rab geranyl geranyl transferase (prenylation of small GTPases of the Rab superfamily); ESCRT: endosomal sorting complex responsible for transport.

9.4.2 HPS

Whereas CHS remains largely mysterious, studies on HPS genes and their corresponding mouse models have greatly accelerated our understanding of melanosome formation and the role of novel trafﬁcking regulators in endosomal dynamics [169]. HPS is a multisystem disorder that is characterized by partial albinism, excessive bleeding, and often lung ﬁbrosis, sometimes accompanied by immunodeﬁciency and/or granulomatous colitis [89]. These systemic symptoms result from the malformation or malfunction of LROs, including melanosomes, platelet dense granules, and, in some cases, lung lamellar bodies, cytotoxic T cell and natural killer cell granules, and perhaps other organelles. HPS results from mutations in any of at least eight different genes; mutations in at least 15 different genes, including the orthologs of the genes that are mutated in human HPS, result in a similar disorder in mice, ﬁrst identiﬁed based on coat-color dilution [89, 170, 171]. Although HPS-associated genes are ubiquitously expressed, they appear to be functionally essential for the generation of a restricted number of LROs such as platelet-dense granules and melanosomes [87]. This cell-type speciﬁcity in penetrance suggests that an ubiquitous set of proteins, perhaps by interacting with additional tissue-speciﬁc machineries, is exploited in specialized cell types to generate a novel lineage of organelles. Understanding how these proteins function in LRO-generating cells such as the melanocyte has already provided insight into novel membrane trafﬁcking pathways that underlie endosomal dynamics and melanosomal enzyme trafﬁcking. 9.4.2.1 Adaptor Protein (AP) Complexes The transfer of macromolecules between organelles of the endosomal system is mediated by tubulovesicular carriers that bud from one membrane and fuse with

272

9 Biogenesis of Melanosomes

another. One critical step in this process is the concentration of proteins into emerging transport carriers through the recognition of sorting signals present in the cytoplasmic domain of transmembrane cargo proteins. Heterotetrameric APs are a family of macromolecular protein complexes that mediate this sorting event. Four distinct APs (AP-1 to -4) are ubiquitously expressed in mammals and each consists of four subunits: one small (σ1–4), one medium (μ1–4), and two large (α, γ, δ, and ε, and β1–4) [172]. The tissue-speciﬁc expression of some “replacement” subunits, such as μ1B and μ3B, leads to the formation of additional unique APs in certain cell types, such as AP-1B and AP-3B in epithelia and neurons, respectively. AP-2 is thought to function exclusively in endocytosis from the plasma membrane by coupling cargo sorting to clathrin-coated vesicle formation. By contrast, AP-1, AP-3, and AP-4 function intracellularly in protein sorting among endosomal organelles and the TGN. AP-3 was the ﬁrst adaptor shown to be involved in melanogenesis. The gene encoding the AP-3 β3A subunit is mutated in HPS2 patients [173] and its mouse model, pearl [174], and the AP-3 δ subunit is mutated in the mouse HPS model, mocha [175]. The mocha and pearl mice are mildly hypopigmented compared to wild-type mice, reﬂecting a diminution in mature melanosomes within melanocytes [176, 177]. AP-3 interacts with di-leucine-based melanosomal sorting signals in the cytoplasmic domains of tyrosinase [28, 40] and OCA2 [44]. Melanocytes derived from HPS2 patients or from pearl mice accumulate tyrosinase in endosomes [28, 178], suggesting a function for AP-3 in sorting tyrosinase from endosomes toward melanosomes. Accordingly, in melanocytes (as in other cell types [179, 180]), the bulk of AP-3 in wild-type melanocytes localizes to clathrin-coated vesicles and buds that are associated with tubular domains of endosomes, and that are often enriched in tyrosinase [28]. While its function in tyrosinase sorting is indisputable, several observations indicate that AP-3 is not required for the transport of other melanosomal enzymes or even for the entire cohort of tyrosinase. (i) Hypopigmentation of pearl and mocha mice is mild compared with other HPS models [169]. (ii) AP-3-deﬁcient melanocytes are still able to generate morphologically intact melanosomes [28, 176, 178] that harbor a reduced but signiﬁcant cohort of tyrosinase [28]. (iii) Other melanosomal proteins, including Tyrp1 and the copper transporter ATP7A, do not interact with AP-3, and their transport to melanosomes is not substantially impaired in AP-3-deﬁcient melanocytes [28, 65, 178] (although misrouting of Tyrp1 through the cell surface seems to be enhanced [181]). Thus, additional sorting molecules must function in melanosome biogenesis. Another critical member of the AP family in melanocytes is AP-1. Genes encoding AP-1 subunits have not been shown to be targets of HPS, but this is likely due to embryonic lethality as observed in mice with targeted deletion in the genes encoding the γ-adaptin [182] or μ1A [183] subunits. The cytoplasmic sorting signals of tyrosinase, Tyrp1, and OCA2 interact with AP-1 [28, 44], and both Tyrp1 and tyrosinase can be detected in vesicles coated with AP-1 in melanocytes [5, 28]. Using in vitro assays, AP-1 and AP-3 have been proposed to mediate the budding of distinct cargo-containing carriers destined for melanosomes [184]. In

9.4 Melanosome Maturation: Cargo Sorting to Mature Melanosomes

melanocytes, AP-1 has been shown to mainly localize to tubulovesicular endosomes that are closely apposed to melanosomes and that derive from the endosomal recycling system; these endosomes likely correspond at least in part to the tubules that contact melanosomes as observed by electron tomography [162]. Whereas in most cells, recycling endosomes accumulate near the microtubule organizing center in the perinuclear region [68], similar domains in melanocytes are positioned in the periphery, apposed to melanosomes. This peripheral polarization of recycling endosomal domains requires AP-1, and facilitates a “dialog” between endosomes and melanosomes that ultimately results in the transfer of protein cargoes, such as Tyrp1, from endosomes to melanosomes[162]. This conclusion is supported by the entrapment of Tyrp1 in endosomes upon small interfering RNA-mediated depletion of AP-1 or an interacting motor protein, KIF13A, in a melanocytic cell line [162, 185]. Together, the data suggest that AP-1 functions both as a cargo sorting molecule and as an effector of endosome positioning to facilitate cargo transfer from recycling endosomal intermediates toward melanosomes. 9.4.2.2 BLOC Complexes In most cell types, AP-1-dependent cargoes are delivered to early endosomes or the TGN and AP-3-dependent cargoes are delivered to late endosomes or lysosomes. By contrast, even though the same compartments are present in melanocytes, a speciﬁc cohort of AP-1- and AP-3-dependent cargoes are efﬁciently delivered to a distinct destination – maturing melanosomes. This suggests that additional effectors must participate in diverting melanosomal cargoes toward this destination. Studies in melanocytes from mouse HPS models have established the biogenesis of lysosome-related organelle complexes (BLOCs) as key effectors in these pathways. The BLOCs are peripheral membrane multisubunit protein complexes that are ubiquitously expressed but, like AP-3, play specialized roles in cells harboring LROs. Genes encoding BLOC subunits constitute the majority of targets of mutation in human and mouse HPS models. There are three different BLOCs: BLOC-1, -2, and -3. Proteins that comprise these multisubunit complexes lack common structural motifs or homology to proteins of known functions [186] and so their molecular functions are not understood. 9.4.2.2.1 BLOC-1 BLOC-1 is an approximately 250-kDa protein complex that consists of eight subunits. Two of the subunits (dysbindin and BLOS3) are encoded by genes that are mutated in human HPS (HPS type 7 and type 8, respectively), and these subunits and three others (Muted, Pallidin, and Cappuccino) are encoded by genes that bear inactivating mutations in mouse HPS models (sandy, reduced pigmentation, muted, pallid, and cappuccino) [170]. Inactivating mutations in any subunit (except BLOS3 in reduced pigmentation mice) destabilize the entire complex, resulting in a complete loss of BLOC-1. BLOC-1 mutant mice are the most severely hypopigmented of all of the mouse HPS models [89, 170] and correspondingly

273

274

9 Biogenesis of Melanosomes

melanocytes derived from them are nearly completely depleted of pigmented melanosomes [176, 177, 187]. By contrast to the substantial missorting of tyrosinase in AP-3-deﬁcient cells, tyrosinase in BLOC-1-deﬁcient melanocytes is only mildly depleted from ﬁbrous stage II-like melanosomes and only partially missorted to endosomes. However, other proteins, including Tyrp1, are nearly completely excluded from melanosomes in BLOC-1-deﬁcient cells and trapped in a continual recycling loop through enlarged vacuolar endosomes [187]; under some conditions, a cohort of Tyrp1 is also misrouted to lysosomes ([181] and our unpublished results). This suggests that BLOC-1 regulates a cargo delivery pathway distinct from that of AP-3. Strikingly, although a substantial cohort of tyrosinase is present in melanosomes in BLOC-1-deﬁcient melanocytes, it is inactive due to an absence of its cofactor, copper [65]. Copper must be pumped into melanosomes by the copper transporter, ATP7A, which itself is a cargo of the BLOC-1dependent transport pathway as shown by its exclusion from melanosomes in BLOC-1-deﬁcient melanocytes; incubation of ﬁxed BLOC-1-deﬁcient cells with excess copper restores tyrosinase activity [65]. Thus, the severe hypopigmentation observed in BLOC-1 mutant mice results from impaired trafﬁcking of Tyrp1, ATP7A, and perhaps other cargoes from endosomes toward melanosomes in an AP-3-independent manner [65, 187]. Like APs, BLOC-1 localizes to early endosomes [181], but consistent with their distinct functions, AP-3 and BLOC-1 are spatially segregated on endosomal membranes: AP-3 is enriched in buds, whereas BLOC-1 localizes to tubular endosomal domains [181], perhaps similar to those that connect vacuolar endosomes with melanosomes [162]. What molecular function BLOC-1 serves on endosomal membrane is still not clear. Analyses of endosomal recycling and localization of Tyrp1 in BLOC-1- and AP-3-deﬁcient melanocytes suggest that BLOC-1 is required for cargo exit from endosomal intermediates, but it remains to be established whether BLOC-1, like AP-1 and AP-3, interacts directly with cargo. BLOC-1 and some of its component subunits have been shown to interact directly with the recycling endosomal tSNARE component syntaxin-13 [188, 189] and the neuronal tSNARE component SNAP-25 [189, 190], suggesting a potential role in regulating fusion steps during melanosomal cargo delivery. BLOC-1 also shows genetic interactions with the recycling endosomal GTPase, Rab11 [189], perhaps suggesting a role in regulating recycling endosomal membrane dynamics. This would be consistent with the role of AP-1 in both recycling endosome dynamics and cargo transport; indeed, the morphological and protein-sorting defects in AP-1-depleted melanocytes are similar to those of BLOC-1-deﬁcient cells [162]. Nevertheless, BLOC-1 has not yet been shown to interact physically with AP-1. Moreover, despite the distinct functions and localization in melanocytes, a cohort of BLOC-1 can be coimmunoprecipitated with AP-3, but not AP-1, from membrane fractions of various cell types [181] and from a subpopulation of synaptic-like microvesicles in neuronal cells [191]. The details of this interaction are still not clear, but one plausible hypothesis is that BLOC-1 and AP-3 might reciprocally regulate their association with membranes [181], and consequently regulate trafﬁcking steps sequentially from the same organelle [181, 191].

9.4 Melanosome Maturation: Cargo Sorting to Mature Melanosomes

9.4.2.2.2 BLOC-2 AP-3, AP-1, and BLOC-1 regulate the sorting of melanosomal cargoes into transport carriers, but what effectors regulate the delivery of these carriers to melanosomes? BLOC-2 might participate in this process. BLOC-2 is a 340-kDa complex that consists of at least three subunits: HPS3, HPS5, and HPS6 [192, 193]. Each subunit is encoded by a gene that is mutated in HPS patients (HPS types 3, 5, and 6) and corresponding mouse models (cocoa, ruby-eye2, and ruby-eye, respectively) [89, 170]. The only recognizable motif on any BLOC-2 subunit is a clathrinbinding domain on HPS3; HPS3 can coimmunoprecipitate with clathrin from melanocyte cell lysates [194], but whether this interaction is physiologically relevant is not clear. BLOC-2-deﬁcient mice are more mildly hypopigmented than BLOC-1 mutants, and defects in melanosome architecture vary from dramatic in the RPE and choroid to less severe in hair-bulb melanocytes [176, 177]. The subcellular distribution of several melanosomal proteins is altered in melanocytes from BLOC-2-deﬁcient patients and mouse models [187, 195–198], but as in BLOC-1deﬁcient cells, Tyrp1 is particularly depleted from melanosomes. Both Tyrp1 and tyrosinase accumulate in vesicles [195, 197] and in large tubulovesicular endosomal structures that are distinct from those that accumulate in BLOC-1 mutants [187], perhaps corresponding to transport intermediates between endosomes and melanosomes [187]. Moreover, Tyrp1 is more rapidly degraded in BLOC-2-deﬁcient melanocytes than in controls [181, 197], suggesting increased missorting to lysosomes. Together, these phenotypes suggest that BLOC-2 might function in the same pathway as BLOC-1 but downstream, perhaps by regulating the directed targeting or fusion of BLOC-1-dependent transport carriers with maturing melanosomes. Consistently, BLOC-2 can be detected on endosomal tubules and can physically interact with a cohort of BLOC-1 [181]. Further analysis is needed to determine whether BLOC-2 indeed functions at this or a distinct step in cargo transport to melanosomes. 9.4.2.2.3 BLOC-3 Although most HPS-associated protein complexes clearly function in vesicular trafﬁcking of melanogenic enzymes, the function of BLOC-3 is less clear. BLOC-3 is an approximately 146-kDa complex consisting of two subunits, HPS1 and HPS4 [199–202], and is largely present in the cytosol with a small fraction associated with membranes [201, 202]. The genes encoding these subunits are mutated in HPS types 1 and 4 – the most prevalent forms of HPS – and corresponding mouse models pale ear and light ear [89, 170]. Melanosomes in RPE and melanocytes of BLOC-3-deﬁcient patients and mice are often misshapen – either grossly enlarged or excessively small and deformed, depending on the tissue [196, 203–206] – but cargo localization defects have not yet been observed [196]. Melanocytes from patients with HPS type 1 display increased autophagy and accumulate melanosomes in compartments with features of autophagic vacuoles [207]. This suggests that the morphological disruption of melanosomes may be a consequence of a defect in a process other than trafﬁcking of melanogenic enzymes. Indeed, in nonpigmented cells, BLOC-3 has been reported to regulate late endosome/

275

276

9 Biogenesis of Melanosomes

lysosome motility and distribution [208], which may have secondary consequences on endosomal membrane dynamics. HPS1 has been localized to the melanosomal membrane and to tubulovesicular membranes close to the Golgi [209]. Interestingly, the only identiﬁed interacting partner of BLOC-3 is the small GTPase Rab9 [200], which functions in recycling from late endosomes/lysosomes. However, despite the GTP dependence of this interaction, suggesting that BLOC-3 is a Rab9 effector [200], it is not clear whether Rab9 functions in melanogenesis. 9.4.3 Molecular Motors and the Cytoskeleton

In melanocytes, fully pigmented stage IV melanosomes accumulate towards the cellular periphery after capture and immobilization on peripheral actin ﬁlaments, ensuring their efﬁcient transfer to keratinocytes (see Chapter 10). However, during their formation and maturation, they undergo rapid bidirectional movements along microtubules, both toward the cell center and toward the cell periphery, as well as shorter localized movements along actin ﬁlaments [210]. These movements are coordinated by molecular motors – myosins on actin, and kinesins and dynein on microtubules. Recent studies suggest that one microtubule motor, the kinesin KIF13A, plays an important role in melanosome maturation. KIF13A is an ubiquitously-expressed plus end-directed kinesin belonging to the kinesin-3 superfamily [211]. KIF13A has been shown to facilitate the transport of mannose 6-phosphate receptors from the TGN to the plasma membrane by virtue of an interaction with the ear domain of the β1-adaptin subunit of AP-1 [211]. KIF13A was identiﬁed in a proteomic analysis of melanosomes [43]. We have shown that in melanocytic cells, KIF13A localizes largely to recycling endosomes and coordinates endosomal positioning in the cell periphery near melanosomes [162]. Consistent with the physical interaction between KIF13A and AP-1, KIF13A depletion from melanocytic cells by small interfering RNA mimics AP-1 depletion by inducing hypopigmentation, perinuclear clustering of recycling endosomes, and entrapment of Tyrp1 on enlarged endosomal vacuoles. This interaction between an AP complex and a molecular motor provides a previously unappreciated link between cargo sorting and endosome positioning to create specialized subdomains of the endosomal recycling system. These specialized domains can be used as gated conduits through which cell-type speciﬁc cargoes are delivered to maturing melanosomes. Our observations suggest that AP-1 and KIF13A might cooperate with AP-3, BLOC-1, and BLOC-2 to control the sorting of melanosomal cargoes from early endosomes and their transport to maturing melanosomes. Deﬁning how these complexes are coordinated during cargo transfer is a challenge for the future. Actin-based motility plays important roles in organelle maturation and fusion events as well. The actin-based motor, myosin Ib, has been proposed to facilitate maturation of stage I melanosomes [212]. Myosin Ib is a monomeric and nonprocessive motor, and has been detected in association with endosomes and the plasma membrane [212]. In melanocytic cells, myosin Ib was shown to coimmu-

9.4 Melanosome Maturation: Cargo Sorting to Mature Melanosomes

noprecipitate with Pmel17, and overexpression of a dominant-negative form of myosin Ib inhibited Pmel17 proteolytic maturation and incorporation in intraluminal vesicles [212]. It is possible that myosin Ib functions to tether Pmel17 to endosomal subdomains destined for invagination to form intralumenal vesicles. Alternatively, it might anchor endosomal subdomains to the actin cytoskeleton to allow for force generation sufﬁcient to drive membrane deformations that are required for intralumenal vesicle formation. In either case, myosin Ib is a candidate to control the formation of stage I melanosomes. 9.4.4 SNAREs, Rabs, and Other Regulators

The directed targeting of AP-3-, AP-1-, and/or BLOC-1-dependent cargo transport carriers toward maturing melanosomes and their fusion with these compartments is likely to require a large cohort of additional molecular components. Membrane fusion processes throughout the secretory and endocytic pathways are mediated by SNARE family proteins [213]. Distinct sets of SNAREs localize to each membrane compartment and transport intermediate, and can be classiﬁed as either vesicle or vSNAREs or components of target or tSNAREs. Fusion occurs when the α-helical regions of cognate vSNAREs on the vesicle/source membrane and tSNARE complexes on the target membrane assemble into a four-helix bundle, resulting in close apposition and destabilization of the two membranes that ultimately results in fusion. To date, the SNAREs that mediate fusion of transport intermediates with maturing melanosomes have not yet been deﬁned. However, multiple endosomal SNAREs are upregulated upon differentiation of a melanocytic cell line [214]. One of them, syntaxin-13, is a strong candidate for facilitating the fusion of recycling endosomal transport intermediates with maturing melanosomes. Although syntaxin-13 has been proposed to function in homotypic endosome–endosome fusion [215], it is predominantly localized to tubular recycling endosomes in many cell types [216] and likely functions in fusion events in the endocytic recycling pathway. Syntaxin-13 has been shown to directly interact in vitro with BLOC-1 via the pallidin subunit [188, 189, 217] and is mislocalized in AP-3-deﬁcient melanocytes [187]. Another excellent candidate is the vesicle-associated membrane protein, VAMP7 (also called TIVAMP for tetanus-insensitive vesicle-associated membrane protein). VAMP7 physically and functionally interacts with AP-3 [218], and is mislocalized and selectively degraded in neuronal cells that lack AP-3 or BLOC-1 [191], suggesting that it might be a key target of these HPS-associated protein complexes. It is tempting to speculate that VAMP7 and syntaxin-13 are components of the SNARE complex that mediates fusion of recycling endosomal tubules with maturing melanosomes. Another likely SNARE regulator is the homotypic fusion and vacuole protein sorting (HOPS) complex. HOPS is an evolutionary conserved complex that appears to function both in membrane tethering and in regulating SNARE-dependent fusion in yeast during homotypic vacuole fusion [219] and likely plays a similar

277

278

9 Biogenesis of Melanosomes

role to regulate endosomal dynamics in mammals [220]. HOPS is comprised of six different subunits. One of them, Vps33, belongs to a family of proteins that interact directly with SNAREs [221], and facilitates SNARE pairing and fusogenic activity [222]. At least two Vps33 isoforms are present in higher eukaryotes and an amino acid substitution in one of them – Vps33a – underlies the HPS-like phenotype of buff mice [223]. Vps33a binds to a number of endosomal SNAREs in mammals, including syntaxin-13 [220], and is therefore poised to regulate fusion between endosomal transport carriers and maturing melanosomes either by recruiting and/or activating SNAREs or by participating directly in the fusion process itself [224]. Melanosomes in RPE of buff mice are small and sparse, and melanocytes derived from buff mice are severely hypopigmented [223], but hair-bulb melanocytes in buff mice show minor defects in melanosome architecture [177]. The Rab family of small GTPases are also key regulators for many membrane trafﬁcking events, particularly in modulating the function and homeostasis of membrane domains [75, 225]. Over 60 different Rabs in mammals function on distinct membranes as molecular switches: when bound to GTP they recruit effectors to their resident membranes, but they release these effectors upon GTP hydrolysis. A number of Rab proteins are selectively expressed at high levels in melanocytes as compared to other tissues [226]. Among them, Rab32 and Rab38 have been shown to be required for the efﬁcient delivery of tyrosinase and Tyrp1 to melanosomes. Rab32 and Rab38 are highly homologous proteins that are expressed in a limited set of cell types, including melanocytes, and that localize in melanocytes to mature melanosomes and tubulovesicular structures that likely correspond to endosomal domains [227]. The Rab38 gene is mutated in ruby rats [228], which suffer from a HPS-like phenotype, and in chocolate mice, which are hypopigmented [229]. Moreover, a mutation in lightoid, the Drosophila melanogaster ortholog of both Rab32 and Rab38, causes defects in eye pigmentation [230]. Interestingly, whereas melanocytes derived from mutant chocolate mice are only mildly hypopigmented, additional depletion of Rab32 severely impairs the delivery of Tyrp1 and tyrosinase to melanosomes, melanosome morphology and pigmentation [227]. These Rab proteins likely function in a redundant way in melanocytes, although perhaps less redundant in RPE [14], to regulate the stability and/or targeting of melanogenic cargo transport intermediates [14]. Additional evidence for the importance of Rabs during melanogenesis comes from analysis of the gunmetal mouse, another of the mouse HPS models. The phenotype results from a mutation in the gene encoding the α subunit of Rab geranylgeranyl transferase II [231], an enzyme that catalyzes prenylation of Rab proteins, which is required for their membrane anchoring and stability. The membrane recruitment of a number of Rabs was greatly reduced in platelets and melanocytes from gunmetal mice relative to controls [226, 231], suggesting that some of them might participate in melanosome biogenesis or function. One was Rab27a, which functions in melanosome motility (see Chapter 10). Another was Rab11, which regulates membrane dynamics within the endocytic recycling

9.5 Conclusions

system [75]. Rab11 was recently predicted to be a BLOC-1-interacting partner [232], and a genetic interaction between Rab11 and BLOC-1 has been observed in D. melanogaster [189]. It is tempting to speculate that Rab11 might regulate BLOC-1dependent cargo delivery between recycling endosomes and melanosomes [162]. The ubiquitously expressed Rab7, which functions in late endosome membrane dynamics and fusion with lysosomes [75], has been implicated in regulating Tyrp1 trafﬁcking [233], but it is not clear whether this reﬂects a direct role for Rab7 in regulating the dynamics or formation of transport intermediates or a more direct role in premelanosome motility [234] or early endosome maturation [235]. Proteomics analyzes show that a number of other Rab proteins associate with melanosomes [42, 43], but whether any of them play functional roles in melanosome formation or motility is not yet clear. 9.4.5 Lipids

One other contributing factor to melanosomal enzyme sorting is the lipid environment of the late secretory and endocytic pathway. Glycosphingolipids are essential components of these membranes, and accumulate in “lipid rafts” at the plasma membrane and in endosomal compartments. Two studies have shown that melanocytes that lack a key enzyme in glycosphingolipid synthesis, ceramide glucosyltransferase, are severely hypopigmented due to missorting of tyrosinase and other melanogenic proteins such as Tyrp1 [236, 237]. Whereas Tyrp1 and tyrosinase in wild-type melanocytes are directly transported from the TGN to endosomal transport intermediates for delivery to melanosomes [156, 237], both proteins are diverted through the plasma membrane in glycosphingolipid-deﬁcient melanocytes [236, 237]. These effects appear to be mediated via the lumenal domains of the melanosomal proteins [236], suggesting a potential interaction between glycosphingolipid head groups and melanosomal enzymes. How these effects are mediated is not yet clear, but represent exciting implications for the role of the lipid bilayer in controlling protein trafﬁcking fates.

9.5 Conclusions

The ﬁndings from the past recent years have started to unravel how melanocytes integrate unique and ubiquitous molecular mechanisms in exploiting the endosomal system to generate melanosomes. The subversion of the endocytic pathway to form melanosomes in melanocytes parallels similar processes for the generation of other LROs, but also additional specialized organelles such as synaptic vesicles in neurons that also originate from specialized endosomal trafﬁcking. In melanocytes, as in other cells hosting LROs, such as platelets, B lymphocytes, and dendritic cells, the multivesicular endosome is a common intermediate playing a key role in the concentration of cargo involved in organelle morphogenesis and

279

280

9 Biogenesis of Melanosomes

function. In melanocytes, these MVBs offer an optimal environment for the formation of Pmel17-driven amyloid ﬁbrils with features of pathological amyloid. The mechanisms, membrane domains, and molecular components involved in the endosomal sorting and the subsequent cleavage of the protein are still far from being fully elucidated. Further studies will help to unravel how within the endosomal membrane this amyloidogenic protein segregates together with proteases, what are the fates of the two domains (amyloidogenic lumenal domain and the C-terminal fragment), and what are the major effectors of these processes. A better comprehension of these mechanisms operating for a physiological amyloid is also certainly an asset to understand the molecular events leading to the formation of pathological amyloid ﬁbrils and their intermediates accumulating in neurodegenerative diseases (Alzheimer’s, prion diseases). In late stages of melanogenesis pigment synthesis is under the control of HPS gene products. Some of these molecular components have been highlighted as novel trafﬁcking regulators. It is unclear how these molecular complexes (BLOC) operate to associate with membranes and regulate cargo sorting. Some of their major interactors have started to be pinpointed mostly from studies in Drosophila and plants (SNAREs, Rab GTPases, retromer components). Additional studies are certainly required to decipher how these ubiquitous components operate in concert with the endosomal trafﬁcking machinery for the generation of tubulovesicular carriers and their transfer to target membranes (i.e., the melanosome). We also need to know more about how melanosome membrane transporters modulate membrane dynamics via the lumenal environment of source and target compartments. Most of the current models for melanosome biogenesis arise from studies on melanocytic cell models of eumelanogenesis. Suitable cell models will be invaluable to enable the elucidation of the basic mechanisms involved in the biogenesis of pheomelanosomes, which will allow conceptual working models to be drawn in parallel for both types of melanogenesis. Finally, given the importance of the keratinocyte in driving melanogenesis, it will be of interest to understand how establishment of the pigmentation synapse inﬂuences melanosome biogenesis and melanosome maturation before ﬁnal transfer.

Acknowledgments

We thank all members of our laboratories for their contributions to many of the studies described here, including Figures 9.1, 9.4, and 9.5 (I. Hurbain), Figure 9.4 (G. van Niel), and Figure 9.5 (D. Tenza). We are grateful for all the daily stimulating discussions. We apologize to those authors whom we did not cite. This work was funded by grants R01 EY015625 and R01 AR048155 from the National Institutes of Health (to M.S.M. and G.R.), and Institut Curie, CNRS, Fondation pour la Recherche Médicale, Institut National du cancer (INCA), Association pour la Recherche sur le Cancer (to G.R.).

References

References 1 King, R.A., Hearing, V.J., Creel, D.J.,

2

3

4

5

6

7

8

9

10

and Oetting, W.S. (1995) Albinism, in The Metabolic and Molecular Bases of Inherited Disease, vol. III (eds C.R. Scriver, A.L. Beaudet, W.S. Sly, and D. Valle), McGraw-Hill, New York, pp. 4353–4392. Marmorstein, A.D., Finnemann, S.C., Bonilha, V.L., and Rodriguez-Boulan, E. (1998) Morphogenesis of the retinal pigment epithelium: toward understanding retinal degenerative diseases. Ann. NY Acad. Sci., 857, 1–12. Carr, R.E. and Siegel, I.M. (1979) The retinal pigment epithelium in ocular albinism, in The Retinal Pigment Epithelium (eds K.M. Zinn and M.P. Marmar), Harvard University Press, Cambridge, MA, pp. 413–423. Orlow, S.J. (1995) Melanosomes are specialized members of the lysosomal lineage of organelles. J. Invest. Dermatol., 105, 3–7. Raposo, G., Tenza, D., Murphy, D.M., Berson, J.F., and Marks, M.S. (2001) Distinct protein sorting and localization to premelanosomes, melanosomes, and lysosomes in pigmented melanocytic cells. J. Cell Biol., 152, 809–824. Marks, M.S. and Seabra, M.C. (2001) The melanosome: membrane dynamics in black and white. Nat. Rev. Mol. Cell Biol., 2, 738–748. Raposo, G. and Marks, M.S. (2007) Melanosomes – dark organelles enlighten endosomal membrane transport. Nat. Rev. Mol. Cell Biol., 8, 786–797. Huizing, M., Helip-Wooley, A., Westbroek, W., Gunay-Aygun, M., and Gahl, W.A. (2008) Disorders of lysosome-related organelle biogenesis: clinical and molecular genetics. Annu. Rev. Genomics Hum. Genet., 9, 359–386. Birbeck, M.S.C., Mercer, E.H., and Barnicot, N.A. (1956) The structure and formation of pigment granules in human hair. Exp. Cell Res., 10, 505–514. Seiji, M., Fitzpatrick, T.M., Simpson, R.T., and Birbeck, M.S.C. (1963) Chemical composition and terminology of specialized organelles (melanosomes

11

12

13

14

15

16

17

18

and melanin granules) in mammalian melanocytes. Nature, 197, 1082–1084. Hurbain, I., Geerts, W.J., Boudier, T., Marco, S., Verkleij, A.J., Marks, M.S., and Raposo, G. (2008) Electron tomography of early melanosomes: implications for melanogenesis and the generation of ﬁbrillar amyloid sheets. Proc. Natl. Acad. Sci. USA, 10519726-31, 19726–19731. Moyer, F.H. (1966) Genetic variations in the ﬁne structure and ontogeny of mouse melanin granules. Am. Zool., 6, 43–66. Lamoreux, M.L., Delmas, V., Larue, L., and Bennett, D.C. (2010) The Colors of Mice: A Model Genetic Network, Wiley-Blackwell, Oxford. Lopes, V.S., Wasmeier, C., Seabra, M.C., and Futter, C.E. (2007) Melanosome maturation defect in Rab38-deﬁcient retinal pigment epithelium results in instability of immature melanosomes during transient melanogenesis. Mol. Biol. Cell, 18, 3914–3927. Miyamoto, M. and Fitzpatrick, T.B. (1957) On the nature of the pigment in retinal pigment epithelium. Science, 126, 449–450. Smith-Thomas, L., Richardson, P., Thody, A.J., Graham, A., Palmer, I., Flemming, L., Parsons, M.A., Rennie, I.G., and MacNeil, S. (1996) Human ocular melanocytes and retinal pigment epithelial cells differ in their melanogenic properties in vivo and in vitro. Curr. Eye Res., 15, 1079–1091. Schraermeyer, U., Kopitz, J., Peters, S., Henke-Fahle, S., Blitgen-Heinecke, P., Kokkinou, D., Schwarz, T., and Bartz-Schmidt, K.U. (2006) Tyrosinase biosynthesis in adult mammalian retinal pigment epithelial cells. Exp. Eye Res., 83, 315–321. Gwynn, B., Ciciotte, S.L., Hunter, S.J., Washburn, L.L., Smith, R.S., Andersen, S.G., Swank, R.T., Dell’Angelica, E.C., Bonifacino, J.S., Eicher, E.M., et al. (2000) Defects in the cappuccino (cno) gene on mouse chromosome 5 and human 4p cause Hermansky–Pudlak

281

282

9 Biogenesis of Melanosomes

19

20

21

22

23

24

25

26

27

28

syndrome by an AP-3-independent mechanism. Blood, 96, 4227–4235. Biesemeier, A., Kreppel, F., Kochanek, S., and Schraermeyer, U. (2010) The classical pathway of melanogenesis is not essential for melanin synthesis in the adult retinal pigment epithelium. Cell Tissue Res., 339, 551–560. Novikoff, A.B., Albala, A., and Biempica, L. (1968) Ultrastructural and cytochemical observations on B-16 and Harding-Passey mouse melanomas. The origin of premelanosomes and compound melanosomes. J. Histochem. Cytochem., 16, 299–319. Seiji, M. and Kikuchi, A. (1969) Acid phosphatase activity in melanosomes. J. Invest. Dermatol., 52, 212–216. Zhou, B.-K., Boissy, R.E., Pifko-Hirst, S., Moran, D.J., and Orlow, S.J. (1993) Lysosome-associated membrane protein-1 (LAMP-1) is the melanocyte vesicular membrane glycoprotein band II. J. Invest. Dermatol., 100, 110–114. Diment, S., Eidelman, M., Rodriguez, G.M., and Orlow, S.J. (1995) Lysosomal hydrolases are present in melanosomes and are elevated in melanizing cells. J. Biol. Chem., 270, 4213–4215. Bennett, D.C. and Lamoreux, M.L. (2003) The color loci of mice – a genetic century. Pigment Cell Res., 16, 333–344. Theos, A.C., Truschel, S.T., Raposo, G., and Marks, M.S. (2005) The Silver locus product Pmel17/gp100/Silv/ME20: controversial in name and in function. Pigment Cell Res., 18, 322–336. Ganesan, A.K., Ho, H., Bodemann, B., Petersen, S., Aruri, J., Koshy, S., Richardson, Z., Le, L.Q., Krasieva, T., Roth, M.G., et al. (2008) Genome-wide siRNA-based functional genomics of pigmentation identiﬁes novel genes and pathways that impact melanogenesis in human cells. PLoS Genet., 4, e1000298. Kushimoto, T., Basrur, V., Valencia, J., Matsunaga, J., Vieira, W.D., Ferrans, V.J., Muller, J., Appella, E., and Hearing, V.J. (2001) A model for melanosome biogenesis based on the puriﬁcation and analysis of early melanosomes. Proc. Natl. Acad. Sci. USA, 98, 10698–10703. Theos, A.C., Tenza, D., Martina, J.A., Hurbain, I., Peden, A.A., Sviderskaya,

29

30

31

32

33

34

35

36

37

38

E.V., Stewart, A., Robinson, M.S., Bennett, D.C., Cutler, D.F., et al. (2005) Functions of AP-3 and AP-1 in tyrosinase sorting from endosomes to melanosomes. Mol. Biol. Cell, 16, 5356–5372. Lerner, A.B., Fitzpatrick, T.B., Calkins, E., and Summerson, W.H. (1949) Mammalian tyrosinase: preparation and properties. J. Biol. Chem., 178, 185–195. Körner, A. and Pawelek, J. (1982) Mammalian tyrosinase catalyzes three reactions in the biosynthesis of melanin. Science, 217, 1163–1165. Medigeshi, G.R. and Schu, P. (2003) Characterization of the in vitro retrograde transport of MPR46. Trafﬁc, 4, 802–811. Solano, F., Jimenez-Cervantes, C., Martinez-Liarte, J.H., Garcia-Borron, J.C., Jara, J.R., and Lozano, J.A. (1996) Molecular mechanism for catalysis by a new zinc-enzyme, dopachrome tautomerase. Biochem. J., 313, 447–453. Kobayashi, T., Urabe, K., Winder, A., Jiménez-Cervantes, C., Imokawa, G., Brewington, T., Solano, F., GarcíaBorrón, J.C., and Hearing, V.J. (1994) Tyrosinase related protein 1 (TRP1) functions as a DHICA oxidase in melanin biosynthesis. EMBO J., 13, 5818–5825. Sarangarajan, R. and Boissy, R.E. (2001) Tyrp1 and oculocutaneous albinism type 3. Pigment Cell Res., 14, 437–444. Kobayashi, T., Imokawa, G., Bennett, D.C., and Hearing, V.J. (1998) Tyrosinase stabilization by Tyrp1 (the brown locus protein). J. Biol. Chem., 273, 31801–31805. Hearing, V.J. (1999) Biochemical control of melanogenesis and melanosomal organization. J. Invest. Dermatol. Symp. Proc., 4, 24–28. Blagoveshchenskaya, A.D., Hewitt, E.W., and Cutler, D.F. (1999) Di-leucine signals mediate targeting of tyrosinase and synaptotagmin to synaptic-like microvesicles within PC12 cells. Mol. Biol. Cell, 10, 3979–3990. Calvo, P.A., Frank, D.W., Bieler, B.M., Berson, J.F., and Marks, M.S. (1999) A cytoplasmic sequence in human tyrosinase deﬁnes a second class of

References

39

40

41

42

43

44

45

46

di-leucine-based sorting signals for late endosomal and lysosomal delivery. J. Biol. Chem., 274, 12780–12789. Vijayasaradhi, S., Xu, Y., Bouchard, B., and Houghton, A.N. (1995) Intracellular sorting and targeting of melanosomal membrane proteins: identiﬁcation of signals for sorting of the human brown locus protein, gp75. J. Cell Biol., 130, 807–820. Honing, S., Sandoval, I.V., and von Figura, K. (1998) A di-leucine-based motif in the cytoplasmic tail of LIMP-II and tyrosinase mediates selective binding of AP-3. EMBO J., 17, 1304–1314. Simmen, T., Schmidt, A., Hunziker, W., and Beermann, F. (1999) The tyrosinase tail mediates sorting to the lysosomal compartment in MDCK cells via a di-leucine and a tyrosine-based signal. J. Cell Sci., 112, 45–53. Basrur, V., Yang, F., Kushimoto, T., Higashimoto, Y., Yasumoto, K., Valencia, J., Muller, J., Vieira, W.D., Watabe, H., Shabanowitz, J., et al. (2003) Proteomic analysis of early melanosomes: identiﬁcation of novel melanosomal proteins. J. Proteome Res., 2, 69–79. Chi, A., Valencia, J.C., Hu, Z.Z., Watabe, H., Yamaguchi, H., Mangini, N.J., Huang, H., Canﬁeld, V.A., Cheng, K.C., Yang, F., et al. (2006) Proteomic and bioinformatic characterization of the biogenesis and function of melanosomes. J. Proteome Res., 5, 3135–3144. Sitaram, A., Piccirillo, R., Palmisano, I., Harper, D.C., Dell’Angelica, E.C., Schiafﬁno, M.V., and Marks, M.C. (2009) Localization to mature melanosomes by virtue of cytoplasmic dileucine motifs is required for human OCA2 function. Mol. Biol. Cell, 29, 1464–1477. Innamorati, G., Piccirillo, R., Bagnato, P., Palmisano, I., and Schiafﬁno, M.V. (2006) The melanosomal/lysosomal protein OA1 has properties of a G protein-coupled receptor. Pigment Cell Res., 19, 125–135. Rinchik, E.M., Bultman, S.J., Horsthemke, B., Lee, S.T., Strunk, K.M.,

47

48

49

50

51

52

53

54

55

Spritz, R.A., Avidano, K.M., Jong, M.T., and Nicholls, R.D. (1993) A gene for the mouse pink-eyed dilution locus and for human type II oculocutaneous albinism. Nature, 361, 72–76. Orlow, S.J. and Brilliant, M.H. (1999) The pink-eyed dilution locus controls the biogenesis of melanosomes and levels of melanosomal proteins in the eye. Exp. Eye Res., 68, 147–154. Rosemblat, S., Sviderskaya, E.V., Easty, D.J., Wilson, A., Kwon, B.S., Bennett, D.C., and Orlow, S.J. (1998) Melanosomal defects in melanocytes from mice lacking expression of the pink-eyed dilution gene: correction by culture in the presence of excess tyrosine. Exp. Cell Res., 239, 344–352. Chen, K., Manga, P., and Orlow, S.J. (2002) Pink-eyed dilution protein controls the processing of tyrosinase. Mol. Biol. Cell, 13, 1953–1964. Costin, G.-E., Valencia, J.C., Vieira, W.D., Lamoreux, M.L., and Hearing, V.J. (2003) Tyrosinase processing and intracellular trafﬁcking is disrupted in mouse primary melanocytes carrying the underwhite (uw) mutation. A model for oculocutaneous albinism (OCA) type 4. J. Cell Sci., 116, 3203–3212. Puri, N., Gardner, J.M., and Brilliant, M.H. (2000) Aberrant pH of melanosomes in pink-eyed dilution (p) mutant melanocytes. J. Invest. Dermatol., 115, 607–613. Brilliant, M. and Gardner, J. (2001) Melanosomal pH, pink locus protein and their roles in melanogenesis. J. Invest. Dermatol., 117, 386–387. Chen, K., Minwalla, L., Ni, L., and Orlow, S.J. (2004) Correction of defective early tyrosinase processing by baﬁlomycin A1 and monensin in pink-eyed dilution melanocytes. Pigment Cell Res., 17, 36–42. Manga, P., Boissy, R.E., Pifko-Hirst, S., Zhou, B.K., and Orlow, S.J. (2001) Mislocalization of melanosomal proteins in melanocytes from mice with oculocutaneous albinism type 2. Exp. Eye Res., 72, 695–710. Schiafﬁno, M.V., Baschirotto, C., Pellegrini, G., Montalti, S., Tacchetti, C., De Luca, M., and Ballabio, A. (1996) The

283

284

9 Biogenesis of Melanosomes

56

57

58

59

60

61

62

63

ocular albinism type 1 gene product is a membrane glycoprotein localized to melanosomes. Proc. Natl. Acad. Sci. USA, 93, 9055–9060. Lopez, V.M., Decatur, C.L., Stamer, W.D., Lynch, R.M., and McKay, B.S. (2008) l-DOPA is an endogenous ligand for OA1. PLoS Biol., 6, e236. Giordano, F., Bonetti, C., Surace, E.M., Marigo, V., and Raposo, G. (2009) The ocular albinism type 1 (OA1) G-protein coupled receptor functions with MART-1 at early stages of melanogenesis to control melanosome identity and composition. Hum. Mol. Genet., 18, 4530–4545. Ginger, R.S., Askew, S.E., Ogborne, R.M., Wilson, S., Ferdinando, D., Dadd, T., Smith, A.M., Kazi, S., Szerencsei, R.T., Winkfein, R.J., et al. (2008) SLC24A5 encodes a trans-Golgi network protein with potassium-dependent sodium-calcium exchange activity that regulates human epidermal melanogenesis. J. Biol. Chem., 283, 5486–5495. Brilliant, M. (2005) Oculocutaneous Albinism Type 4, GeneReviews, Seattle, WA. Lamason, R.L., Mohideen, M.-A.P.K., Mest, J.R., Wong, A.C., Norton, H.L., Aros, M.C., Jurynec, M.J., Mao, X., Humphreville, V.R., Humbert, J.E., et al. (2005) SLC24A5, a putative cation exchanger, affects pigmentation in zebraﬁsh and humans. Science, 310, 1782–1786. Newton, J.M., Cohen-Barak, O., Hagiwara, N., Gardner, J.M., Davisson, M.T., King, R.A., and Brilliant, M.H. (2001) Mutations in the human orthologue of the mouse underwhite gene (uw) underlie a new form of oculocutaneous albinism, OCA4. Am. J. Hum. Genet., 69, 981–988. Oancea, E., Vriens, J., Brauchi, S., Jun, J., Splawski, I., and Clapham, D.E. (2009) TRPM1 forms ion channels associated with melanin content in melanocytes. Sci. Signal., 2, ra21. McNeill, M.S., Paulsen, J., Bonde, G., Burnight, E., Hsu, M.Y., and Cornell, R.A. (2007) Cell death of melanophores in zebraﬁsh trpm7 mutant embryos

64

65

66

67

68

69

70

71

72

73

74 75

depends on melanin synthesis. J. Invest. Dermatol., 127, 2020–2030. Hoashi, T., Watabe, H., Muller, J., Yamaguchi, Y., Vieira, W.D., and Hearing, V.J. (2005) MART-1 is required for the function of the melanosomal matrix protein PMEL17/GP100 and the maturation of melanosomes. J. Biol. Chem., 280, 14006–14016. Setty, S.R., Tenza, D., Sviderskaya, E.V., Bennett, D.C., Raposo, G., and Marks, M.S. (2008) Cell-speciﬁc ATP7A transport sustains copper-dependent tyrosinase activity in melanosomes. Nature, 454, 1142–1146. Lutsenko, S., Barnes, N.L., Bartee, M.Y., and Dmitriev, O.Y. (2007) Function and regulation of human coppertransporting ATPases. Physiol. Rev., 87, 1011–1046. Silvers, W.K. (1979) The Coat Colors of Mice: A Model for Mammalian Gene Action and Interaction, Springer, New York. Maxﬁeld, F.R. and McGraw, T.E. (2004) Endocytic recycling. Nat. Rev. Mol. Cell Biol., 5, 121–132. Gruenberg, J. (2001) The endocytic pathway: a mosaic of domains. Nat. Rev. Mol. Cell Biol., 2, 721–730. Gould, G.W. and Lippincott-Schwartz, J. (2009) New roles for endosomes: from vesicular carriers to multi-purpose platforms. Nat. Rev. Mol. Cell Biol., 10, 287–292. Doherty, G.J. and McMahon, H.T. (2009) Mechanisms of endocytosis. Annu. Rev. Biochem., 78, 857–902. Jovic, M., Sharma, M., Rahajeng, J., and Caplan, S. (2010) The early endosome: a busy sorting station for proteins at the crossroads. Histol. Histopathol., 25, 99–112. Simonsen, A., Lippe, R., Christoforidis, S., Gaullier, J.M., Brech, A., Callaghan, J., Toh, B.H., Murphy, C., Zerial, M., and Stenmark, H. (1998) EEA1 links PI3K function to Rab5 regulation of endosome fusion. Nature, 394, 494–498. van Ijzendoorn, S.C. (2006) Recycling endosomes. J. Cell Sci., 119, 1679–1681. Stenmark, H. (2009) Rab GTPases as coordinators of vesicle trafﬁc. Nat. Rev. Mol. Cell., 10, 513–525.

References 76 Piper, R.C. and Katzmann, D.J. (2007)

77

78

79

80

81

82

83

84

85

86

87

Biogenesis and function of multivesicular bodies. Annu. Rev. Cell Dev. Biol., 23, 519–547. Sachse, M., Urbe, S., Oorschot, V., Strous, G.J., and Klumperman, J. (2002) Bilayered clathrin coats on endosomal vacuoles are involved in protein sorting toward lysosomes. Mol. Biol. Cell, 13, 1313–1328. Raiborg, C. and Stenmark, H. (2009) The ESCRT machinery in endosomal sorting of ubiquitylated membrane proteins. Nature, 458, 445–452. Hurley, J.H. (2010) The ESCRT complexes. Crit. Rev. Biochem. Mol. Biol., 45, 463–487. Pryor, P.R. and Luzio, J.P. (2009) Delivery of endocytosed membrane proteins to the lysosome. Biochim. Biophys. Acta, 1793, 615–624. White, I.J., Bailey, L.M., Aghakhani, M.R., Moss, S.E., and Futter, C.E. (2006) EGF stimulates annexin 1-dependent inward vesiculation in a multivesicular endosome subpopulation. EMBO J., 25, 1–12. Simons, M. and Raposo, G. (2009) Exosomes – vesicular carriers for intercellular communication. Curr. Opin. Cell Biol., 21, 575–581. Turner, W.A., Jr, Taylor, J.D., and Tchen, T.T. (1975) Melanosome formation in the goldﬁsh: the role of multivesicular bodies. J. Ultrastruct. Res., 51, 16–31. Jimbow, K., Oikawa, O., Sugiyama, S., and Takeuchi, T. (1979) Comparison of eumelanogenesis in retinal and follicular melanocytes; role of vesiculo-globular bodies in melanosome differentiation. J. Invest. Dermatol., 73, 278–284. Maul, G.G. and Brumbaugh, J.A. (1971) On the possible function of coated vesicles in melanogenesis of the regenerating fowl feather. J. Cell Biol., 48, 41–48. Bouchard, B., Fuller, B.B., Vijayasaradhi, S., and Houghton, A.N. (1989) Induction of pigmentation in mouse ﬁbroblasts by expression of human tyrosinase cDNA. J. Exp. Med., 169, 2029–2042. Raposo, G., Marks, M.S., and Cutler, D.F. (2007) Lysosome-related organelles:

88

89

90

91

92

93

94

95

96

97

driving post-Golgi compartments into specialisation. Curr. Opin. Cell Biol., 19, 394–401. Stinchcombe, J., Bossi, G., and Grifﬁths, G.M. (2004) Linking albinism and immunity: the secrets of secretory lysosomes. Science, 305, 55–59. Wei, M.L. (2006) Hermansky–Pudlak syndrome: a disease of protein trafﬁcking and organelle function. Pigment Cell Res., 19, 19–42. Boissy, R.E., Moellmann, G.E., and Halaban, R. (1987) Tyrosinase and acid phosphatase activities in melanocytes from avian albinos. J. Invest. Dermatol., 88, 292–300. Anderson, R.G.W., Falck, J.R., Goldstein, J.L., and Brown, M.S. (1984) Visualization of acidic organelles in intact cells by electron microscopy. Proc. Natl. Acad. Sci. USA, 81, 4838–4842. Fujita, H., Sasano, E., Yasunaga, K., Furuta, K., Yokota, S., Wada, I., and Himeno, M. (2001) Evidence for distinct membrane trafﬁc pathways to melanosomes and lysosomes in melanocytes. J. Investig. Dermatol. Symp. Proc., 6, 19–24. Schraermeyer, U. (1995) Transport of endocytosed material into melanin granules in cultured choroidal melanocytes of cattle – new insights into the relationship of melanosomes with lysosomes. Pigment Cell Res., 8, 209–214. Orlow, S.J., Boissy, R.E., Moran, D.J., and Pifko-Hirst, S. (1993) Subcellular distribution of tyrosinase and tyrosinaserelated protein-1: implications for melanosomal biogenesis. J. Invest. Dermatol., 100, 55–64. Seiji, M. and Iwashia, S. (1965) Intracellular localization of tyrosinase and site of melanin formation in melanocyte. J. Invest. Dermatol., 45, 305–314. Saeki, H. and Oikawa, A. (1978) Effects of pH and type of sugar in the medium on tyrosinase activity in cultured melanoma cells. J. Cell Physiol., 94, 139–145. Devi, C.C., Tripathi, R.K., and Ramaiah, A. (1987) pH-dependent interconvertible allosteric forms of murine melanoma

285

286

9 Biogenesis of Melanosomes

98

99

100

101

102

103

104

105

106

tyrosinase. Physiological implications. Eur. J. Biochem., 166, 705–711. Berson, J.F., Harper, D., Tenza, D., Raposo, G., and Marks, M.S. (2001) Pmel17 initiates premelanosome morphogenesis within multivesicular bodies. Mol. Biol. Cell, 12, 3451–3464. Piccirillo, R., Palmisano, I., Innamorati, G., Bagnato, P., Altimare, D., and Schiafﬁno, M.V. (2006) An unconventional dileucine-based motif and a novel cytosolic motif are required for the lysosomal and melanosomal targeting of OA1. J. Cell Sci., 119, 2003–2014. Furumura, M., Sakai, C., Potterf, S.B., Vieira, W.D., Barsh, G.S., and Hearing, V.J. (1998) Characterization of genes modulated during pheomelanogenesis using differential display. Proc. Natl. Acad. Sci. USA, 95, 7374–7378. Kobayashi, T., Vieira, W.D., Potterf, B., Sakai, C., Imokawa, G., and Hearing, V.J. (1995) Modulation of melanogenic protein expression during the switch from eu- to pheomelanogenesis. J. Cell Sci., 108, 2301–2309. Schraermeyer, U. and Heimann, K. (1999) Current understanding on the role of retinal pigment epithelium and its pigmentation. Pigment Cell Res., 12, 219–236. Schraermeyer, U., Peters, S., Thumann, G., Kociok, N., and Heimann, K. (1999) Melanin granules of retinal pigment epithelium are connected with the lysosomal degradation pathway. Exp. Eye Res., 68, 237–245. Nichols, S.E., Harper, D.C., Berson, J.F., and Marks, M.S. (2003) A novel splice variant of Pmel17 expressed by human melanocytes and melanoma cells lacking some of the internal repeats. J. Invest. Dermatol., 121, 821–830. Bailin, T., Lee, S.T., and Spritz, R.A. (1996) Genomic organization and sequence of D12S53E (Pmel 17), the human homologue of the mouse silver (si) locus. J. Invest. Dermatol., 106, 24–27. Chakraborty, A.K., Platt, J.T., Kim, K.K., Kwon, B.S., Bennett, D.C., and Pawelek, J.M. (1996) Polymerization of 5,6-dihydroxyindole-2-carboxylic acid to

107

108

109

110

111

112

113

melanin by the pmel 17/silver locus protein. Eur. J. Biochem., 236, 180–188. Adema, G.J., de Boer, A.J., Vogel, A.M., Loenen, W.A., and Figdor, C.G. (1994) Molecular characterization of the melanocyte lineage-speciﬁc antigen gp100. J. Biol. Chem., 269, 20126–20133. Hoashi, T., Muller, J., Vieira, W.D., Rouzaud, F., Kikuchi, K., Tamaki, K., and Hearing, V.J. (2006) The repeat domain of the melanosomal matrix protein PMEL17/GP100 is required for the formation of organellar ﬁbers. J. Biol. Chem., 281, 21198–21208. Hoashi, T., Sato, S., Yamaguchi, Y., Passeron, T., Tamaki, K., and Hearing, V.J. (2010) Glycoprotein nonmetastatic melanoma protein b, a melanocytic cell marker, is a melanosome-speciﬁc and proteolytically released protein. FASEB J., 24, 1616–1629. Wagner, S.N., Wagner, C., Schultewolter, T., and Goos, M. (1997) Analysis of Pmel17/gp100 expression in primary human tissue specimens: implications for melanoma immunoand gene-therapy. Cancer Immunol. Immunother., 44, 239–247. Cormier, J.N., Abati, A., Fetsch, P., Hijazi, Y.M., Rosenberg, S.A., Marincola, F.M., and Topalian, S.L. (1998) Comparative analysis of the in vivo expression of tyrosinase, MART-1/ Melan-A, and gp100 in metastatic melanoma lesions: implications for immunotherapy. J. Immunother., 21, 27–31. Folpe, A.L., Goodman, Z.D., Ishak, K.G., Paulino, A.F., Taboada, E.M., Meehan, S.A., and Weiss, S.W. (2000) Clear cell myomelanocytic tumor of the falciform ligament/ligamentum teres: a novel member of the perivascular epithelioid clear cell family of tumors with a predilection for children and young adults. Am. J. Surg. Pathol., 24, 1239–1246. Matsumoto, Y., Horiba, K., Usuki, J., Chu, S.C., Ferrans, V.J., and Moss, J. (1999) Markers of cell proliferation and expression of melanosomal antigen in lymphangioleiomyomatosis. Am. J. Respir. Cell Mol. Biol., 21, 327–336.

References 114 Lee, Z.H., Hou, L., Moellmann, G.,

115

116

117

118

119

120

121

Kuklinska, E., Antol, K., Fraser, M., Halaban, R., and Kwon, B.S. (1996) Characterization and subcellular localization of human Pmel 17/silver, a 100-kDa (pre)melanosomal membrane protein associated with 5,6-dihydroxyindole-2-carboxylic acid (DHICA) converting activity. J. Invest. Dermatol., 106, 605–610. Orlow, S.J., Zhou, B.-K., Boissy, R.E., and Pifko-Hirst, S. (1993) Identiﬁcation of a mammalian melanosomal matrix glycoprotein. J. Invest. Dermatol., 101, 141–144. Zhou, B.K., Kobayashi, T., Donatien, P.D., Bennett, D.C., Hearing, V.J., and Orlow, S.J. (1994) Identiﬁcation of a melanosomal matrix protein encoded by the murine si (silver) locus using “organelle scanning”. Proc. Natl. Acad. Sci. USA, 91, 7076–7080. Berson, J.F., Theos, A.C., Harper, D.C., Tenza, D., Raposo, G., and Marks, M.S. (2003) Proprotein convertase cleavage liberates a ﬁbrillogenic fragment of a resident glycoprotein to initiate melanosome biogenesis. J. Cell Biol., 161, 521–533. Donatien, P.D. and Orlow, S.J. (1995) Interaction of melanosomal proteins with melanin. Eur. J. Biochem., 232, 159–164. Theos, A.C., Berson, J.F., Theos, S.C., Herman, K.E., Harper, D.C., Tenza, D., Sviderskaya, E.V., Lamoreux, M.L., Bennett, D.C., Raposo, G., et al. (2006) Dual loss of ER export and endocytic signals with altered melanosome morphology in the silver mutation of Pmel17. Mol. Biol. Cell, 17, 3598–3612. Fowler, D.M., Koulov, A.V., Alory-Jost, C., Marks, M.S., Balch, W.E., and Kelly, J.W. (2006) Functional amyloid formation within mammalian tissue. PLoS Biol., 4, e6. Watt, B., van Niel, G., Fowler, D.M., Hurbain, I., Luk, K.C., Stayrook, S.E., Lemmon, M.A., Raposo, G., Shorter, J., Kelly, J.W., et al. (2009) N-terminal domains elicit formation of functional Pmel17 amyloid ﬁbrils. J. Biol. Chem., 284, 35543–35555.

122 Chiti, F. and Dobson, C.M. (2006)

123

124

125

126

127

128

129

130

Protein misfolding,functional amyloid and human disease. Annu. Rev. Biochem., 75, 333–366. Fowler, D.M., Koulov, A.V., Balch, W.E., and Kelly, J.W. (2007) Functional amyloid-from bacteria to humans. Trends Biochem. Sci., 32, 217–223. Watt, B., Raposo, G., and Marks, M.S. (2010) Pmel17: an amyloid determinant of organelle structure, in Functional Amyloid Aggregation (ed. M. Bucciantini), Research Signpost, Trivandrum. Esclamado, R.M., Gown, A.M., and Vogel, A.M. (1986) Unique proteins deﬁned by monoclonal antibodies speciﬁc for human melanoma. Am. J. Surg., 152, 376–385. Gown, A.M., Vogel, A.M., Hoak, D., Gough, F., and McNutt, M.A. (1986) Monoclonal antibodies speciﬁc for melanocytic tumors distinguish subpopulations of melanocytes. Am. J. Pathol., 123, 195–203. Chiamenti, A.M., Vella, F., Bonetti, F., Pea, M., Ferrari, S., Martignoni, G., Benedetti, A., and Suzuki, H. (1996) Anti-melanoma monoclonal antibody HMB-45 on enhanced chemiluminescence-western blotting recognizes a 30–35 kDa melanosomeassociated sialated glycoprotein. Melanoma Res., 6, 291–298. Kapur, R.P., Bigler, S.A., Skelly, M., and Gown, A.M. (1992) Anti-melanoma monoclonal antibody HMB45 identiﬁes an oncofetal glycoconjugate associated with immature melanosomes. J. Histochem. Cytochem., 40, 207–212. Harper, D.C., Theos, A.C., Herman, K.E., Tenza, D., Raposo, G., and Marks, M.S. (2008) Premelanosome amyloid-like ﬁbrils are composed of only Golgiprocessed forms of pmel17 that have been proteolytically processed in endosomes. J. Biol. Chem., 283, 2307–2322. Lepage, S. and Lapointe, R. (2006) Melanosomal targeting sequences from gp100 are essential for MHC class II–restricted endogenous epitope presentation and mobilization to endosomal compartments. Cancer Res., 66, 2423–2432.

287

288

9 Biogenesis of Melanosomes 131 Robila, V., Ostankovitch, M., Altrich-

132

133

134

135

136

137

138

139

Vanlith, M.L., Theos, A.C., Drover, S., Marks, M.S., Restifo, N., and Engelhard, V.H. (2008) MHC class II presentation of gp100 epitopes in melanoma cells requires the function of conventional endosomes and is inﬂuenced by melanosomes. J. Immunol., 181, 7843–7852. Kummer, M.P., Maruyama, H., Huelsmann, C., Baches, S., Weggen, S., and Koo, E.H. (2009) Formation of pmel17 amyloid is regulated by juxtamembrane metalloproteinase cleavage, and the resulting C-terminal fragment is a substrate for γ-secretase. J. Biol. Chem., 284, 2296–2306. Maresh, G.A., Marken, J.S., Neubauer, M., Aruffo, A., Hellström, I., Hellström, K.E., and Marquardt, H. (1994) Cloning and expression of the gene for the melanoma-associated ME20 antigen. DNA Cell Biol., 13, 87–95. Hoashi, T., Tamaki, K., and Hearing, V.J. (2010) The secreted form of a melanocyte membrane-bound glycoprotein (Pmel17/gp100) is released by ectodomain shedding. FASEB J., 24, 916–930. De Strooper, B., Vassar, R., and Golde, T. (2010) The secretases: enzymes with therapeutic potential in Alzheimer disease. Nat. Rev. Neurol., 6, 99–107. McGlinchey, R.P., Shewmaker, F., McPhie, P., Monterroso, B., Thurber, K., and Wickner, R.B. (2009) The repeat domain of the melanosome ﬁbril protein Pmel17 forms the amyloid core promoting melanin synthesis. Proc. Natl. Acad. Sci. USA, 106, 13731–13736. Futter, C.E., Ramalho, J.S., Jaissle, G.B., Seeliger, M.W., and Seabra, M.C. (2004) The role of Rab27a in the regulation of melanosome distribution within retinal pigment epithelial cells. Mol. Biol. Cell, 15, 2264–2275. Dunn, L.C. and Thigpen, L.W. (1930) The silver mouse: a recessive color variation. J. Hered., 21, 495. Bassi, M.T., Schiafﬁno, M.V., Renieri, A., De Nigris, F., Galli, L., Bruttini, M., Gebbia, M., Bergen, A.A., Lewis, R.A., and Ballabio, A. (1995) Cloning of the gene for ocular albinism type 1 from the

140

141

142

143

144

145

146

147

distal short arm of the X chromosome. Nat. Genet., 10, 13–19. Schiafﬁno, M.V., d’Addio, M., Alloni, A., Baschirotto, C., Valetti, C., Cortese, K., Puri, C., Bassi, M.T., Colla, C., De Luca, M., et al. (1999) Ocular albinism: evidence for a defect in an intracellular signal transduction system. Nat. Genet., 23, 108–112. Cortese, K., Giordano, F., Surace, E.M., Venturi, C., Ballabio, A., Tacchetti, C., and Marigo, V. (2005) The ocular albinism type 1 (OA1) gene controls melanosome maturation and size. Invest. Ophthalmol. Vis. Sci., 46, 4358–4364. Incerti, B., Cortese, K., Pizzigoni, A., Surace, E.M., Varani, S., Coppola, M., Jeffery, G., Seeliger, M., Jaissle, G., Bennett, D.C., et al. (2000) Oa1 knock-out: new insights on the pathogenesis of ocular albinism type 1. Hum. Mol. Genet., 9, 2781–2788. Young, A., Powelson, E.B., Whitney, I.E., Raven, M.A., Nusinowitz, S., Jiang, M., Birnbaumer, L., Reese, B.E., and Farber, D.B. (2008) Involvement of OA1, an intracellular GPCR, and G alpha i3, its binding protein, in melanosomal biogenesis and optic pathway formation. Invest. Ophthalmol. Vis. Sci., 49, 3245–3252. Blander, J.M. (2007) Coupling Toll-like receptor signaling with phagocytosis: potentiation of antigen presentation. Trends Immunol., 28, 19–25. Palmisano, I., Bagnato, P., Palmigiano, A., Innamorati, G., Rotondo, G., Altimare, D., Venturi, C., Sviderskaya, E.V., Piccirillo, R., Coppola, M., et al. (2008) The ocular albinism type 1 protein, an intracellular G proteincoupled receptor, regulates melanosome transport in pigment cells. Hum. Mol. Genet., 17, 3487–3501. Schiafﬁno, M.V. (2010) Signaling pathways in melanosome biogenesis and pathology. Int. J. Biochem. Cell Biol., 42, 1094–1104. Theos, A.C., Truschel, S.T., Tenza, D., Hurbain, I., Harper, D.C., Berson, J.F., Thomas, P.C., Raposo, G., and Marks, M.S. (2006) A lumenal domaindependent pathway for sorting to intralumenal vesicles of multivesicular

References

148

149

150

151

152

153

154

155

endosomes involved in organelle morphogenesis. Dev. Cell, 10, 343–354. Kovacs, G.G., Gelpi, E., Ströbel, T., Ricken, G., Nyengaard, J.R., Bernheimer, H., and Budka, H. (2007) Involvement of the endosomal– lysosomal system correlates with regional pathology in Creutzfeldt–Jakob disease. J. Neuropathol. Exp. Neurol., 66, 628–636. Maul, G.G. (1969) Golgi–melanosome relationship in human melanoma in vitro. J. Ultrastruct. Res., 26, 163–176. Kobayashi, T., Urabe, K., Orlow, S.J., Higashi, K., Imokawa, G., Kwon, B.S., Potterf, B., and Hearing, V.J. (1994) The Pmel 17/silver locus protein. Characterization and investigation of its melanogenic function. J. Biol. Chem., 269, 29198–29205. Yasumoto, K., Watabe, H., Valencia, J.C., Kushimoto, T., Kobayashi, T., Appella, E., and Hearing, V.J. (2004) Epitope mapping of the melanosomal matrix protein gp100 (PMEL17): rapid processing in the endoplasmic reticulum and glycosylation in the early Golgi compartment. J. Biol. Chem., 279, 28330–28338. Valencia, J.C., Rouzaud, F., Julien, S., Chen, K.G., Passeron, T., Yamaguchi, Y., Abu-Asab, M., Tsokos, M., Costin, G.E., Yamaguchi, H., et al. (2007) Sialylated core 1 O-glycans inﬂuence the sorting of Pmel17/gp100 and determine its capacity to form ﬁbrils. J. Biol. Chem., 282, 11266–11280. Valencia, J.C., Hoashi, T., Pawelek, J.M., Solano, F., and Hearing, V.J. (2006) Pmel17: controversial indeed but critical to melanocyte function. Pigment Cell Res., 19, 250–252; author reply 253–257. Gagnon, E., Duclos, S., Rondeau, C., Chevet, E., Cameron, P.H., SteeleMortimer, O., Paiement, J., Bergeron, J.J., and Desjardins, M. (2002) Endoplasmic reticulum-mediated phagocytosis is a mechanism of entry into macrophages. Cell, 110, 119–131. Lévy, F., Muehlethaler, K., Salvi, S., Peitrequin, A.-L., Lindholm, C.K., Cerottini, J.-C., and Rimoldi, D. (2005) Ubiquitylation of a melanosomal protein by HECT-E3 ligases serves as sorting

156

157

158

159

160

161

162

163

164

signal for lysosomal degradation. Mol. Biol. Cell, 16, 1777–1787. Truschel, S.T., Simoes, S., Setty, S.R.G., Harper, D.C., Tenza, T., Thomas, P.C., Herman, K.E., Sackett, S.D., Cowan, D.C., Theos, A.C., et al. (2009) ESCRT-I function is required for Tyrp-1 transport from early endosomes to the melanosome limiting membrane. Trafﬁc, 10, 1318–1336. Trajkovic, K., Hsu, C., Chiantia, S., Rajendran, L., Wenzel, D., Wieland, F., Schwille, P., Brugger, B., and Simons, M. (2008) Ceramide triggers budding of exosome vesicles into multivesicular endosomes. Science, 319, 1244–1247. Arvan, P. and Castle, D. (1998) Sorting and storage during secretory granule biogenesis: looking backward and looking forward. Biochem. J., 332, 593–610. Gibson, A., Futter, C.E., Maxwell, S., Allchin, E.H., Shipman, M., Kraehenbuhl, J.P., Domingo, D., Odorizzi, G., Trowbridge, I.S., and Hopkins, C.R. (1998) Sorting mechanisms regulating membrane protein trafﬁc in the apical transcytotic pathway of polarized MDCK cells. J. Cell Biol., 143, 81–94. Gruenberg, J. and Maxﬁeld, F.R. (1995) Membrane transport in the endocytic pathway. Curr. Opin. Cell Biol., 7, 552–563. Bright, N.A., Gratian, M.J., and Luzio, J.P. (2005) Endocytic delivery to lysosomes mediated by concurrent fusion and kissing events in living cells. Curr. Biol., 15, 360–365. Delevoye, C., Hurbain, I., Tenza, D., Sibarita, J.-B., Uzan-Gafsou, S., Ohno, H., Geerts, W.J.C., Verkleij, A.J., Salamero, J., Marks, M.S., et al. (2009) AP-1 and KIF13A coordinate endosomal sorting and positioning during melanosome biogenesis. J. Cell Biol., 187, 247–264. Seabra, M.C. and Coudrier, E. (2004) Rab GTPases and myosin motors in organelle motility. Trafﬁc, 5, 393–399. Van Den Bossche, K., Naeyaert, J.M., and Lambert, J. (2006) The quest for the mechanism of melanin transfer. Trafﬁc, 7, 769–778.

289

290

9 Biogenesis of Melanosomes 165 Perou, C.M., Moore, K.J., Nagle, D.L.,

166

167

168

169

170

171

172

173

Misumi, D.J., Woolf, E.A., McGrail, S.H., Holmgren, L., Brody, T.H., Dussault, B.J., Jr, Monroe, C.A., et al. (1996) Identiﬁcation of the murine beige gene by YAC complementation and positional cloning. Nat. Genet., 13, 303–308. Barbosa, M.D., Nguyen, Q.A., Tchernev, V.T., Ashley, J.A., Detter, J.C., Blaydes, S.M., Brandt, S.J., Chotai, D., Hodgman, C., Solari, R.C., et al. (1996) Identiﬁcation of the homologous beige and Chediak–Higashi syndrome genes. Nature, 382, 262–265. Ward, D.M., Shiﬂett, S.L., Huynh, D., Vaughn, M.B., Prestwich, G., and Kaplan, J. (2003) Use of expression constructs to dissect the functional domains of the CHS/beige protein: identiﬁcation of multiple phenotypes. Trafﬁc, 4, 403–415. Faigle, W., Raposo, G., Tenza, D., Pinet, V., Vogt, A.B., Kropshofer, H., Fischer, A., de Saint-Basile, G., and Amigorena, S. (1998) Deﬁcient peptide loading and MHC class II endosomal sorting in a human genetic immunodeﬁciency disease: the Chediak–Higashi syndrome. J. Cell Biol., 141, 1121–1134. Gautam, R., Novak, E.K., Tan, J., Wakamatsu, K., Ito, S., and Swank, R.T. (2006) Interaction of Hermansky–Pudlak syndrome genes in the regulation of lysosome-related organelles. Trafﬁc, 7, 779–792. Di Pietro, S.M. and Dell’Angelica, E.C. (2005) The cell biology of Hermansky– Pudlak syndrome: recent advances. Trafﬁc, 6, 525–533. Swank, R.T., Novak, E.K., McGarry, M.P., Zhang, Y., Li, W., Zhang, Q., and Feng, L. (2000) Abnormal vesicular trafﬁcking in mouse models of Hermansky–Pudlak syndrome. Pigment Cell Res., 13, 59–67. Ohno, H. (2006) Cell science at a glance: clathrin-associated adaptor protein complexes. J. Cell Sci., 119, 3719–3721. Dell’Angelica, E.C., Shotelersuk, V., Aguilar, R.C., Gahl, W.A., and Bonifacino, J.S. (1999) Altered trafﬁcking of lysosomal proteins in Hermansky–Pudlak syndrome due to

174

175

176

177

178

179

180

181

mutations in the β3A subunit of the AP-3 adaptor. Mol. Cell, 3, 11–21. Feng, L., Seymour, A.B., Jiang, S., To, A., Peden, A.A., Novak, E.K., Zhen, L., Rusiniak, M.E., Eicher, E.M., Robinson, M.S., et al. (1999) The beta3A subunit gene (Ap3b1) of the AP-3 adaptor complex is altered in the mouse hypopigmentation mutant pearl, a model for Hermansky–Pudlak syndrome and night blindness. Hum. Mol. Genet., 8, 323–330. Kantheti, P., Qiao, X., Diaz, M.E., Peden, A.A., Meyer, G.E., Carskadon, S.L., Kapfhamer, D., Sufalko, D., Robinson, M.S., Noebels, J.L., et al. (1998) Mutation in AP-3β in the mocha mouse links endosomal transport to storage deﬁciency in platelets, melanosomes, and synaptic vesicles. Neuron, 21, 111–122. Nguyen, T., Novak, E.K., Kermani, M., Fluhr, J., Peters, L.L., Swank, R.T., and Wei, M.L. (2002) Melanosome morphologies in murine models of Hermansky–Pudlak syndrome reﬂect blocks in organelle development. J. Invest. Dermatol., 119, 1156–1164. Nguyen, T. and Wei, M.L. (2004) Characterization of melanosomes in murine Hermansky–Pudlak syndrome: mechanisms of hypopigmentation. J. Invest. Dermatol., 122, 452–460. Huizing, M., Sarangarajan, R., Strovel, E., Zho, Y., Gahl, W.A., and Boissy, R.E. (2001) AP-3 mediates tyrosinase but not TRP-1 trafﬁcking in human melanocytes. Mol. Biol. Cell, 12, 2075–2085. Dell’Angelica, E.C., Klumperman, J., Stoorvogel, W., and Bonifacino, J.S. (1998) Association of the AP-3 adaptor complex with clathrin. Science, 280, 431–434. Peden, A.A., Oorschot, V., Hesser, B.A., Austin, C.D., Scheller, R.H., and Klumperman, J. (2004) Localization of the AP-3 adaptor complex deﬁnes a novel endosomal exit site for lysosomal membrane proteins. J. Cell Biol., 164, 1065–1076. Di Pietro, S.M., Falcon-Perez, J.M., Tenza, D., Setty, S.R., Marks, M.S., Raposo, G., and Dell’Angelica, E.C.

References

182

183

184

185

186

187

188

189

(2006) BLOC-1 interacts with BLOC-2 and the AP-3 complex to facilitate protein trafﬁcking on endosomes. Mol. Biol. Cell, 17, 4027–4038. Zizioli, D., Meyer, C., Guhde, G., Saftig, P., von Figura, K., and Schu, P. (1999) Early embryonic death of mice deﬁcient in gamma-adaptin. J. Biol. Chem., 274, 5385–5390. Meyer, C., Zizioli, D., Lausmann, S., Eskelinen, E.L., Hamann, J., Saftig, P., von Figura, K., and Schu, P. (2000) mu1A-adaptin-deﬁcient mice: lethality, loss of AP-1 binding and rerouting of mannose 6-phosphate receptors. EMBO J., 19, 2193–2203. Chapuy, B., Tikkanen, R., Muhlhausen, C., Wenzel, D., von Figura, K., and Honing, S. (2008) AP-1 and AP-3 mediate sorting of melanosomal and lysosomal membrane proteins into distinct post-Golgi trafﬁcking pathways. Trafﬁc, 9, 1157–1172. Lakkaraju, A., Carvajal-Gonzalez, J.M., and Rodriguez-Boulan, E. (2009) It takes two to tango to the melanosome. J. Cell Biol., 187, 161–163. Dell’Angelica, E.C. (2004) The building BLOC(k)s of lysosomes and related organelles. Curr. Opin. Cell Biol., 16, 458–464. Setty, S.R., Tenza, D., Truschel, S.T., Chou, E., Sviderskaya, E.V., Theos, A.C., Lamoreux, M.L., Di Pietro, S.M., Starcevic, M., Bennett, D.C., et al. (2007) BLOC-1 is required for cargo-speciﬁc sorting from vacuolar early endosomes toward lysosome-related organelles. Mol. Biol. Cell, 18, 768–780. Moriyama, K. and Bonifacino, J.S. (2002) Pallidin is a component of a multiprotein complex involved in the biogenesis of lysosome-related organelles. Trafﬁc, 3, 666–677. Ghiani, C.A., Starcevic, M., RodriguezFernandez, I.A., Nazarian, R., Cheli, V.T., Chan, L.N., Malvar, J.S., de Vellis, J., Sabatti, C., and Dell’Angelica, E.C. (2010) The dysbindin-containing complex (BLOC-1) in brain: developmental regulation, interaction with SNARE proteins and role in neurite outgrowth. Mol. Psychiatry, 15, 204–215.

190 Ilardi, J.M., Mochida, S., and Sheng,

191

192

193

194

195

196

197

Z.H. (1999) Snapin: a SNARE-associated protein implicated in synaptic transmission. Nat. Neurosci., 2, 119–124. Salazar, G., Craige, B., Styers, M.L., Newell-Litwa, K.A., Doucette, M.M., Wainer, B.H., Falcon-Perez, J.M., Dell’Angelica, E.C., Peden, A.A., Werner, E., et al. (2006) BLOC-1 complex deﬁciency alters the targeting of adaptor protein complex-3 cargoes. Mol. Biol. Cell, 14, 4014–4026. Di Pietro, S.M., Falcon-Perez, J.M., and Dell’Angelica, E.C. (2004) Characterization of BLOC-2, a complex containing the Hermansky–Pudlak syndrome proteins HPS3, HPS5 and HPS6. Trafﬁc, 5, 276–283. Gautam, R., Chintala, S., Li, W., Zhang, Q., Tan, J., Novak, E.K., Di Pietro, S.M., Dell’Angelica, E.C., and Swank, R.T. (2004) The Hermansky–Pudlak syndrome 3 (cocoa) protein is a component of the biogenesis of lysosome-related organelles complex-2 (BLOC-2). J. Biol. Chem., 279, 12935–12942. Helip-Wooley, A., Westbroek, W., Dorward, H., Mommaas, M., Boissy, R.E., Gahl, W.A., and Huizing, M. (2005) Association of the Hermansky– Pudlak syndrome type-3 protein with clathrin. BMC Cell Biol., 6, 33. Boissy, R.E., Richmond, B., Huizing, M., Helip-Wooley, A., Zhao, Y., Koshoffer, A., and Gahl, W.A. (2005) Melanocyte-speciﬁc proteins are aberrantly trafﬁcked in melanocytes of Hermansky–Pudlak syndrome-type 3. Am. J. Pathol., 166, 231–240. Richmond, B., Huizing, M., Knapp, J., Koshoffer, A., Zhao, Y., Gahl, W.A., and Boissy, R.E. (2005) Melanocytes derived from patients with Hermansky–Pudlak syndrome types 1, 2, and 3 have distinct defects in cargo trafﬁcking. J. Invest. Dermatol., 124, 420–427. Helip-Wooley, A., Westbroek, W., Dorward, H.M., Koshoffer, A., Huizing, M., Boissy, R.E., and Gahl, W.A. (2007) Improper trafﬁcking of melanocytespeciﬁc proteins in Hermansky–Pudlak syndrome type-5. J. Invest. Dermatol., 127, 1471–1478.

291

292

9 Biogenesis of Melanosomes 198 Huizing, M., Pederson, B., Hess, R.A.,

199

200

201

202

203

204

205

Grifﬁn, A., Helip-Wooley, A., Westbroek, W., Dorward, H., O’Brien, K.J., Golas, G., Tsilou, E., et al. (2009) Clinical and cellular characterisation of Hermansky–Pudlak syndrome type 6. J. Med. Genet., 46, 803–810. Chiang, P.-W., Oiso, N., Gautam, R., Swank, R.T., and Spritz, R.A. (2003) The Hermansky–Pudlak syndrome 1 (HPS1) and HPS4 proteins are components of two complexes, BLOC-3 and BLOC-4, involved in the biogenesis of lysosomerelated organelles. J. Biol. Chem., 278, 20332–20337. Kloer, D.R., Rojas, R., Ivan, V., Moriyama, K., van Vlijmen, T., Murthy, N., Ghirlando, R., van der Sluijs, P., Hurley, J.H., and Bonifacino, J.S. (2010) Assembly of the biogenesis of lysosomerelated organelles complex-3 (BLOC-3) and its interaction with Rab9. J. Biol. Chem., 285, 7794–7804. Martina, J.A., Moriyama, K., and Bonifacino, J.S. (2003) BLOC-3, a protein complex containing the Hermansky–Pudlak syndrome gene products HPS1 and HPS4. J. Biol. Chem., 278, 29376–29384. Nazarian, R., Falcon-Perez, J.M., and Dell’Angelica, E.C. (2003) Biogenesis of lysosome-related organelles complex 3 (BLOC-3): a complex containing the Hermansky–Pudlak syndrome (HPS) proteins HPS1 and HPS4. Proc. Natl. Acad. Sci. USA, 100, 8770–8775. Gardner, J.M., Wildenberg, S.C., Keiper, N.M., Novak, E.K., Rusiniak, M.E., Swank, R.T., Puri, N., Finger, J.N., Hagiwara, N., Lehman, A.L., et al. (1997) The mouse pale ear (ep) mutation is the homologue of human Hermansky– Pudlak syndrome. Proc. Natl. Acad. Sci. USA, 94, 9238–9243. Nguyen, T. and Wei, M.L. (2007) Hermansky–Pudlak HPS1/pale ear gene regulates epidermal and dermal melanocyte development. J. Invest. Dermatol., 127, 421–428. Suzuki, T., Li, W., Zhang, Q., Karim, A., Novak, E.K., Sviderskaya, E.V., Hill, S.P., Bennett, D.C., Levin, A.V., Nieuwenhuis, H.K., et al. (2002) Hermansky–Pudlak syndrome is caused by mutations in

206

207

208

209

210

211

212

213

HPS4, the human homolog of the mouse light-ear gene. Nat. Genet., 30, 321–324. Horikawa, T., Araki, K., Fukai, K., Ueda, M., Ueda, T., Ito, S., and Ichihashi, M. (2000) Heterozygous HPS1 mutations in a case of Hermansky–Pudlak syndrome with giant melanosomes. Br. J. Dermatol., 143, 635–640. Smith, J.W., Koshoffer, A., Morris, R.E., and Boissy, R.E. (2005) Membranous complexes characteristic of melanocytes derived from patients with Hermansky– Pudlak syndrome type 1 are macroautophagosomal entities of the lysosomal compartment. Pigment Cell Res., 18, 417–426. Falcon-Perez, J.M., Nazarian, R., Sabatti, C., and Dell’Angelica, E.C. (2005) Distribution and dynamics of Lamp1containing endocytic organelles in ﬁbroblasts deﬁcient in BLOC-3. J. Cell Sci., 118, 5243–5255. Oh, J., Liu, Z.X., Feng, G.H., Raposo, G., and Spritz, R.A. (2000) The Hermansky–Pudlak syndrome (HPS) protein is part of a high molecular weight complex involved in biogenesis of early melanosomes. Hum. Mol. Genet., 9, 375–385. Wu, X., Bowers, B., Rao, K., Wei, Q., and Hammer, J.A., III (1998) Visualization of melanosome dynamics within wild-type and dilute melanocytes suggests a paradigm for myosin V function in vivo. J. Cell Biol., 143, 1899–1918. Nakagawa, T., Setou, M., Seog, D., Ogasawara, K., Dohmae, N., Takio, K., and Hirokawa, N. (2000) A novel motor, KIF13A, transports mannose-6phosphate receptor to plasma membrane through direct interaction with AP-1 complex. Cell, 103, 569–581. Salas-Cortes, L., Ye, F., Tenza, D., Wilhelm, C., Theos, A., Louvard, D., Raposo, G., and Coudrier, E. (2005) Myosin Ib modulates the morphology and the protein transport within multi-vesicular sorting endosomes. J. Cell Sci., 118, 4823–4832. Bonifacino, J.S. and Glick, B.S. (2004) The mechanisms of vesicle budding and fusion. Cell, 116, 153–166.

References 214 Wade, N., Bryant, N.J., Connolly, L.M.,

215

216

217

218

219

220

221

222

Simpson, R.J., Luzio, J.P., Piper, R.C., and James, D.E. (2001) Syntaxin 7 complexes with mouse Vps10p tail interactor Ib, Syntaxin 6, vesicleassociated membrane protein (VAMP)8, and VAMP7 in B16 melanoma cells. J. Biol. Chem., 276, 19820–19827. McBride, H.M., Rybin, V., Murph, y.C., Giner, A., Teasdale, R., and Zerial, M. (1999) Oligomeric complexes link Rab5 effectors with NSF and drive membrane fusion via interactions between EEA1 and syntaxin 13. Cell, 98, 377–386. Prekeris, R., Klumperman, J., Chen, Y.A., and Scheller, R.H. (1998) Syntaxin 13 mediates cycling of plasma membrane proteins via tubulovesicular recycling endosomes. J. Cell Biol., 143, 957–971. Huang, L., Kuo, Y.M., and Gitschier, J. (1999) The pallid gene encodes a novel, syntaxin 13-interacting protein involved in platelet storage pool deﬁciency. Nat. Genet., 23, 329–332. Martinez-Arca, S., Rudge, R., Vacca, M., Raposo, G., Camonis, J., ProuxGillardeaux, V., Daviet, L., Formstecher, E., Hamburger, A., Filippini, F., et al. (2003) A dual mechanism controlling the localization and function of exocytic v-SNAREs. Proc. Natl. Acad. Sci. USA, 100, 9011–9016. Sato, T.K., Rehling, P., Peterson, M.R., and Emr, S.D. (2000) Class C vps protein complex regulates vacuolar SNARE pairing and is required for vesicle docking/fusion. Mol. Cell, 6, 661–671. Richardson, S.C., Winistorfer, S.C., Poupon, V., Luzio, J.P., and Piper, R.C. (2004) Mammalian late vacuole protein sorting orthologues participate in early endosomal fusion and interact with the cytoskeleton. Mol. Biol. Cell, 15, 1197–1210. Gerst, J.E. (2003) SNARE regulators: matchmakers and matchbreakers. Biochim. Biophys. Acta, 1641, 99–110. Shen, J., Tareste, D.C., Paumet, F., Rothman, J.E., and Melia, T.J. (2007) Selective activation of cognate SNAREpins by Sec1/Munc18 proteins. Cell, 128, 183–195.

223 Suzuki, T., Oiso, N., Gautam, R., Novak,

224

225

226

227

228

229

230

231

E.K., Panthier, J.J., Suprabha, P.G., Vida, T., Swank, R.T., and Spritz, R.A. (2003) The mouse organellar biogenesis mutant buff results from a mutation in Vps33a, a homologue of yeast vps33 and Drosophila carnation. Proc. Natl. Acad. Sci. USA, 21, 21. Nickerson, D.P., Brett, C.L., and Merz, A.J. (2009) Vps-C complexes: gatekeepers of endolysosomal trafﬁc. Curr. Opin. Cell Biol., 21, 543–551. Zerial, M. and McBride, H. (2001) Rab proteins as membrane organizers. Nat. Rev. Mol. Cell Biol., 2, 107–119. Zhang, Q., Zhen, L., Li, W., Novak, E.K., Collinson, L.M., Jang, E.K., Haslam, R.J., Elliott, R.W., and Swank, R.T. (2002) Cell-speciﬁc abnormal prenylation of Rab proteins in platelets and melanocytes of the gunmetal mouse. Br. J. Haematol., 117, 414–423. Wasmeier, C., Romao, M., Plowright, L., Bennett, D.C., Raposo, G., and Seabra, M.C. (2006) Rab38 and Rab32 control post-Golgi trafﬁcking of melanogenic enzymes. J. Cell Biol., 175, 271–281. Oiso, N., Riddle, S.R., Serikawa, T., Kuramoto, T., and Spritz, R.A. (2004) The rat Ruby (R) locus is Rab38: identical mutations in Fawn-hooded and Tester-Moriyama rats derived from an ancestral Long Evans rat sub-strain. Mamm. Genome, 15, 307–314. Loftus, S.K., Larson, D.M., Baxter, L.L., Antonellis, A., Chen, Y., Wu, X., Jiang, Y., Bittner, M., Hammer, J.A., 3rd, and Pavan, W.J. (2002) Mutation of melanosome protein RAB38 in chocolate mice. Proc. Natl. Acad. Sci. USA, 99, 4471–4476. Ma, J., Plesken, H., Treisman, J.E., Edelman-Novemsky, I., and Ren, M. (2004) Lightoid and Claret: a rab GTPase and its putative guanine nucleotide exchange factor in biogenesis of Drosophila eye pigment granules. Proc. Natl. Acad. Sci. USA, 101, 11652–11657. Detter, J.C., Zhang, Q., Mules, E.H., Novak, E.K., Mishra, V.S., Li, W., McMurtrie, E.B., Tchernev, V.T., Wallace, M.R., Seabra, M.C., et al. (2000) Rab geranylgeranyl transferase alpha mutation in the gunmetal mouse

293

294

9 Biogenesis of Melanosomes reduces Rab prenylation and platelet synthesis. Proc. Natl. Acad. Sci. USA, 97, 4144–4149. 232 Rodriguez-Fernandez, I.A. and Dell’Angelica, E.C. (2009) A data-mining approach to rank candidate proteinbinding partners – the case of biogenesis of lysosome-related organelles complex-1 (BLOC-1). J. Inherit. Metab. Dis., 32, 190–203. 233 Hirosaki, K., Yamashita, T., Wada, I., Jin, H.Y., and Jimbow, K. (2002) Tyrosinase and tyrosinase-related protein 1 require Rab7 for their intracellular transport. J. Invest. Dermatol., 119, 475–480. 234 Jordens, I., Westbroek, W., Marsman, M., Rocha, N., Mommaas, M., Huizing, M., Lambert, J., Naeyaert, J.M., and Neefjes, J. (2006) Rab7 and Rab27a control two motor protein activities involved in melanosomal transport. Pigment Cell Res., 19, 412–423.

235 Rink, J., Ghigo, E., Kalaidzidis, Y., and

Zerial, M. (2005) Rab conversion as a mechanism of progression from early to late endosomes. Cell, 122, 735–749. 236 Groux-Degroote, S., van Dijk, S.M., Wolthoorn, J., Neumann, S., Theos, A.C., De Mazière, A.M., Klumperman, J., van Meer, G., and Sprong, H. (2008) Glycolipid-dependent sorting of melanosomal from lysosomal membrane proteins by lumenal determinants. Trafﬁc, 9, 951–963. 237 Sprong, H., Degroote, S., Claessens, T., van Drunen, J., Oorschot, V., Westerink, B.H., Hirabayashi, Y., Klumperman, J., van der Sluijs, P., and van Meer, G. (2001) Glycosphingolipids are required for sorting melanosomal proteins in the Golgi complex. J. Cell Biol., 155, 369–380.

295

10 Transport and Distribution of Melanosomes Mireille Van Gele and Jo Lambert

10.1 Introduction

The light-absorbing pigment melanin is produced and stored within specialized lysosome-related organelles termed melanosomes, present in the melanocytes [1]. Two major types of melanin are synthesized inside melanosomes: eumelanin (black/brown) and pheomelanin (yellow/red). The formation of both pigments starts with the enzymatic oxidation of tyrosine, but then follows different metabolic pathways (see Chapter 4). Each epidermal melanocyte provides melanin to approximately 36 keratinocytes using its dendrites. After transfer, melanin is transported to the apical face of the keratinocyte nucleus, where it protects the genetic material against UV damage. This symbiotic system is called the “epidermal melanin unit” and is primarily responsible for skin pigmentation. Extensive studies on pigment cells of lower vertebrates and mammals revealed that the transport of melanosomes from their site of synthesis (i.e. cell center) to the cell periphery occurs via fast, long-range transport along microtubules, followed by a short-range movement along actin ﬁlaments. Three different classes of motor proteins were shown to be involved in this intracellular transport: kinesins, dyneins, and myosin Va (Myo5a). In melanophores of ﬁsh and frogs they are responsible for fast aggregation and dispersion of melanosomes in response to variations in the environment [2]. In retinal pigment epithelium (RPE) cells of the eye these motor proteins guarantee transport of melanosomes into the apical processes in response to light [3]. Studies of naturally occurring mouse color mutants revealed that the transport of melanosomes along actin ﬁlaments and their capture in the cell periphery is accomplished by the action of a Rab27a–Mlph (melanophilin)– Myo5a tripartite complex. The importance of this complex was also conﬁrmed in human melanocytes and mutations in MYO5A, RAB27A, or MLPH result in Griscelli syndrome (GS) types I, II, and III, respectively. These patients are characterized by pigment dilution of the skin and hair, due to defects involving melanosome transport in melanocytes. A similar tripartite complex, consisting of Rab27a–Myrip–Myo7a (myosin VIIa), was discovered in mouse RPE cells.

Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

296

10 Transport and Distribution of Melanosomes

Likewise, this complex is responsible for the transport of melanosomes along actin ﬁlaments and their tethering in actin-rich apical processes. Epidermal melanocytes transfer mature melanosomes along the dendritic processes to neighboring keratinocytes. The mechanisms behind this transfer process have proved to be difﬁcult to unravel. Different hypotheses, such as exocytosis, cytophagocytosis, and tunnel formation, were proposed. With the development of high-resolution assays (in space as well as in time) the evidence is growing that ﬁlopodia are used as conduits for intercellular transfer of melanosomes. Whether ﬁlopodia eventually connect cytoplasms forming a tunnel or they are phagocytosed by the receiving cell (the keratinocyte), as argued by Singh et al., remains to be further substantiated [4]. Inside keratinocytes the melanosomes continue their journey and travel along microtubules in the direction of the cell center. They tend to aggregate above the keratinocyte nucleus in lightly pigmented skin, while in darker phototypes they are dispersed throughout the cytoplasm. Little is known about the signals that regulate this distribution pattern. Communication and melanin transfer between individual keratinocytes may play an important role in this process.

10.2 Model Systems to Study Pigment Transport 10.2.1 Melanophores from Fish and Amphibians

In ﬁsh and amphibians the melanosomes are found in melanophores – large black pigmented cells present in the dermis and epidermis. The major function of these cells involves aggregation of pigment granules in the center of the cell (resulting in skin lightening) or dispersal of them throughout the cytoplasm (causing a dark appearance of the skin). This bidirectional and coordinated transport of pigment granules (i.e., physiological color change) allows the animal to effect color changes important for camouﬂage and social interactions. Similar to mammals, the melanosomes of amphibians can also be dispersed and transferred to surrounding cells, providing long-term morphological changes. Much of our current understanding of melanosome motility and its regulation is derived from studies of ﬁsh and frog melanophores (Figure 10.1a). They are regarded as excellent model systems for the analysis of organelle transport for several reasons: (i) intracellular movements of their large pigment granules can easily be monitored by conventional bright-ﬁeld microscopy, (ii) bidirectional transport can be experimentally manipulated by treatment of the cells with neurotransmitters (ﬁsh) or hormonal stimuli (ﬁsh and frogs), and (iii) the availability of an immortalized cell line of Xenopus melanophores [7]. In ﬁsh, melanosome movements are fast and tightly regulated due to the longer and unidirectional movements that they undergo. In frog melanophores, melanosome transport is bidirectional, including frequent pauses and reversals that

10.2 Model Systems to Study Pigment Transport a)

b)

c)

Figure 10.1 (a) Bright ﬁeld images of frog melanophores (Xenopus laevis) illustrating

dispersed pigment (left) and aggregated pigment (right). Bars = 10 μm. (Reproduced from [5].) (b) Schematic view of a melanophore showing the dispersal and aggregation of pigment granules that occur along microtubules under the inﬂuence of cAMP. (c) Bright ﬁeld images of wild-type (left) and ashen mouse melanocytes (right). Bars = 12 μm. (Reproduced from [6].)

297

298

10 Transport and Distribution of Melanosomes

increase the amount of time necessary to aggregate or disperse. In this context, transport in frog melanophores resembles more the organelle transport occurring in other cell types. In both cases, however, melanosome dispersion is induced by elevation of intracellular cAMP levels, while aggregation is triggered by depression of cAMP (Figure 10.1b). The downstream steps of these second messengers are poorly understood but seem to involve protein kinases and phosphatases [5]. Several morphological and functional transport studies in melanophores revealed that two elements of the cytoskeleton – microtubules and actin ﬁlaments – are required for the movement of pigment in these cells. Microtubuledependence of melanosome transport could be demonstrated by treating ﬁsh melanophores with low temperature, high hydrostatic pressure, or microtubuledisrupting drugs. These treatments resulted in the inhibition of both aggregation and dispersion [8, 9]. Additional studies showed that dynein, a minus-end-directed microtubule motor, colocalized with ﬁsh [10] and amphibian melanosomes [11], and that it was involved in the aggregation of these melanosomes [12]. By using cultured Xenopus melanophores, Rogers et al. were also able to show that kinesin-2 was present on melanosomes [11]. A year later, Tuma et al. showed, by using a dominant-negative mutant of kinesin-2, that it was also the motor protein responsible for dispersion of melanosomes in Xenopus melanophores [13]. The same authors suggested that kinesin-2, and not conventional kinesin, is responsible for dispersion in ﬁsh melanophores as well [14]. Parallel to the studies described above, other research groups performed studies on ﬁsh melanophores that could, however, demonstrate that removal of microtubules did not completely abolish pigment movement, suggesting also a role for the actin cytoskeleton in pigment transport [15]. In addition, studies on frog melanophores have shown that removal of actin ﬁlaments inhibited dispersion of melanosomes [16, 17] and that actin ﬁlaments were indeed present on the cell periphery of melanophores [18]. Later, it was demonstrated that the motor protein Myo5a was responsible for actin-based transport of melanosomes in Xenopus melanophores [19]. Moreover, the actin ﬁlaments were mainly required for proper dispersion and for keeping the melanophores dispersed [17]. Next, Sköld et al. demonstrated that Myo5a was present on ﬁsh melanosomes as well, but the actinbased transport seemed to play a less important role for the overall bidirectional translocations in ﬁsh [20]. To discover in more detail how the motor proteins dynein, kinesin, and Myo5a are regulated in melanophores and cooperate with each other to move along their cytoskeleton tracks we refer the reader to two excellent reviews on this topic [2, 21]. 10.2.2 Mammalian Melanocytes

Mammalian melanocytes have been widely used to study the transport of melanosomes. Extensive studies of mouse and human melanocytes showed that, similar to what is observed in melanophores of lower vertebrates, both microtubules and actin ﬁlaments are required for melanosome transport. A major advantage of

10.3 Intracellular Melanosome Transport

mammalian melanocytes lies in the fact that a number of naturally occurring mouse coat color mutants with defects in melanosome transport exist: dilute, ashen, and leaden [22–24]. Normal levels of melanin are synthesized in these mouse mutants, but melanosomes are not efﬁciently distributed to neighboring keratinocytes. Instead they clump around the melanocyte cell nucleus (Figure 10.1c). Molecular genetic studies led to the discovery of the genes involved and uncovered the role of the Rab27a–Mlph–Myo5a complex in melanosome transport. Myo5a is the actin-based motor protein that transports melanosomes along the sublemmal and ﬁlopodial network. Two molecules – Mlph and Rab27a – are necessary for the attachment of Myo5a to the melanosome surface [25]. The important role of this tripartite complex in melanosome transport was also conﬁrmed in human melanocytes and the mouse coat color mutants ﬁnd their human counterparts in GS type 1 (MYO5A), 2 (RAB27A), and 3 (MLPH) [26]. 10.2.3 RPE Cells

RPE cells lie at the back of the vertebrate eye, and they synthesize melanin pigments and store them within numerous melanosomes. RPE cells closely interact with photoreceptors – a critical action for the maintenance of visual function. In eyes of lower vertebrates, which do not have dilatable pupils, the melanosomes in the RPE cells undergo massive, light-dependent migrations, whereas the movements in mammals are more subtle. Some of the properties of pigment transport in RPE cells are similar to those observed in melanophores. For instance, the bidirectional transport of melanosomes and the induction of pigment aggregation or dispersion by treatment with cAMP or dopamine, respectively. Fish RPE cells have mainly been used to study the dual role of microtubules and actin ﬁlaments in melanosome transport [27]. Studies on mouse cells led to the discovery that the actin-based motor Myo7a is involved in melanosome transport in RPE cells, whereas Myo5a accomplishes this function in epidermal melanocytes [28]. This ﬁnding provided evidence that Rab27a can link several classes of myosin motors to the organelle surface [3, 29] (see Section 10.4).

10.3 Intracellular Melanosome Transport

Inside mammalian melanocytes, mature melanosomes are transported from the perinuclear area to the cell periphery and dendritic tips along the microtubular network. This fast, long-distance transport is mediated by two classes of cytoskeletal motor proteins: dyneins and kinesins. Upon arrival at the cell periphery, melanosomes undergo “short-range” movements along the sublemmal actin network through the association with the processive motor protein Myo5a, which attaches to melanosomes through interaction with Mlph and Rab27a (Figure 10.2).

299

300

10 Transport and Distribution of Melanosomes

Intracellular transport of melanosomes. The tip of a melanocyte dendrite is depicted. Two motor proteins transport melanosomes along microtubules: kinesin centrifugally and dynein centripetally. Dynein mainly localizes to early melanosomes, and probably binds them via interaction with Rilp and Rab7. Mature melanosomes travel towards the dendrite

Figure 10.2

tips where they hop onto the sublemmal actin network. Transport along the actin ﬁlaments is accomplished by the action of the Rab27a–Mlph–Myo5a complex. When Rab27a binds another effector, Slp2-a, melanosomes attach to the plasma membrane. Melanosomes are now ready for release and transfer to keratinocytes.

10.3.1 Microtubule-Based Transport 10.3.1.1 Kinesin and Dynein Two motor proteins orchestrate the transport of melanosomes along microtubules: kinesin for anterograde movement towards the microtubule plus-ends located at the cell periphery and dynein for retrograde movement towards the microtubule minus-ends located at the cell center [30, 31]. This area is actually less extensively studied in melanocytes and knowledge from other cell systems, such as neurons, opens areas for future research around these motors in melanocytes [32]. Melanocytic dendrites consist of a central core of microtubules and a subcortical actin network. In previous reports we showed the presence of microtubuleassociated motor proteins kinesin and cytoplasmic dynein on the melanosomal surface, forming a link with microtubules [33, 34]. We could also demonstrate the association of kinectin, the kinesin receptor, with melanosomes. The interaction of cytoplasmic dynein with its cargoes is thought to be indirectly mediated by dynactin, a complex that binds to the dynein intermediate chain. The dynactin subunits P150Glued and P50 colocalize with the melanosomal membrane [35]. Spe-

10.3 Intracellular Melanosome Transport

ciﬁcally, Gross et al. propose that dynactin enables dynein to participate efﬁciently in bidirectional transport, increasing its ability to stay “on” during minus-end motion and keeping it “off” during plus-end motion [36]. Dynein is the common motor protein found in early, unpigmented, melanosomes. Probably, Rab7 recruits dynein via its interaction with Rab7-interacting lysosomal protein (Rilp). Rab7 mainly localizes to early stage melanosomes and these melanosomes locate to the cell center upon overexpression of Rilp [37]. Instead, kinesin is enriched in mature melanosomes [38]. This is consistent with the ongoing maturation of early melanosomes in the perinuclear area and the transport of mature melanosomes, ready for delivery to keratinocytes, to the dendrite tips (Figure 10.2). Next to their role in melanosomal organelle transport, these motors also play a role in positioning and correctly localizing endosomal proteins and domains during the biogenesis of melanosomes [38, 39] 10.3.2 Actin-Based Transport 10.3.2.1 MYO5A MYO5A functions as a dimeric, actin-based molecular motor protein, and is divided into three separate domains termed “head”, “neck, ” and “tail” (Figure 10.3). First,

Figure 10.3 Schematic view of active MYO5A. At the N-terminus MYO5A is composed of two motor domains (MD) that contain ATP- and actin-binding sites followed by a neck domain consisting of six IQ motifs. The tail region comprises a proximal and

medial domain, and contains three major α-helical coiled-coil segments interrupted by two small uncoiled regions. The distal or globular tail domain at the C-terminus is responsible for cargo binding via cargospeciﬁc receptors.

301

302

10 Transport and Distribution of Melanosomes

MYO5A is composed out of two separate N-terminal (or “head”) motor domains, containing ATP-binding sites as well as having the capability to bind to actin ﬁbers. These ATP-binding sites can convert energy released by multiple catalytic cycles of ATP hydrolysis, generating a mechanical movement along the actin ﬁlaments. Following these two separate head domains are two identical heavy chains that dimerize into each other. These two α-helical coils termed the “neck” domains are comprised of six subsequent amino acid residues termed IQ motifs with the consensus sequence IQXXXRGXXXR. These IQ motifs act as binding sites for light chains, which are either calmodulin or calmodulin-like light chains and have a regulatory function by controlling the ATPase activity of the globular heads [40]. The “tail” (proximal/medial) domain commences at the site where both α-helical coils dimerize to form a homodimer made up of a series of coiled-coil segments interrupted by small ﬂexible globular regions (around 500 amino acids). The length and composition of the medial tail region varies depending on alternative splicing. In humans, six different isoforms have been identiﬁed that are expressed in certain cell types, and each isoform speciﬁes which cargo is bound and transported. Transcripts containing the exon F were found to be the isoforms intervening in melanosome transport [41]. Finally, the last approximately 400 amino acids distinguish the distal/globular tail region, also known as the cargo-binding site, where the interaction with different cargos is mediated by organelle speciﬁc receptors. Mercer et al. were the ﬁrst to report that the murine Myo5a (dilute) locus encodes the heavy chain of Myo5a, a processive molecular motor [23]. More then 20 years ago, the GS was mapped by Pastural et al. to 15q21, the region where MYO5A is located [42]. The presence of mutations in MYO5A [42] and in its murine counterpart [23] led to the identiﬁcation of MYO5A to be the ﬁrst gene involved in GS. This disease was therefore termed GS1 (OMIM #214450). In general, these patients show hypomelanosis with a primary neurological deﬁcit and without immunological impairment or manifestations of hemophagocytic syndrome. MYO5A mutations were identiﬁed in only two GS patients [42, 43]. However, the low detection frequency of MYO5A mutations in other patients [44] suggested the existence of a second GS gene also located at 15q21 [31, 43]. 10.3.2.2 RAB27A The Ras super family of small GTPase monomeric G-proteins (20–25 kDa) is essential in regulating a wide variety of cell processes, including vesicular transport pathways, cellular differentiation, and motility [45]. RAB27A is a member of this family and has the ability to act as a molecular switch by cycling between two conformations through the binding of GTP, rendering it in an “active” state or hydrolyzing GTP to GDP, translating it to its “inactive” form. Two regions have been shown to change conformation upon GDP or GTP binding and have been termed switch I and switch II regions. A guanine nucleotide exchange factor (GEF), called Rab3GEF, is responsible for the activation of Rab27a in melanocytes [46]. Rab27a is a substrate for prenylation by an enzyme called Rab geranylgeranyl transferase. These prenyl groups are anchored to mature melanosomes and bind

10.3 Intracellular Melanosome Transport

to two conserved cysteine residues located at the C-terminal end, also referred to as geranylgeranylation motifs of Rab27a. This interaction can be considered similar to the labeling or tagging of membranes of vesicles, thus deﬁning the identity and future routing of these vesicles. This implies that Rab27a is involved in targeting, docking and fusion of transport vesicles with their appropriate acceptor membranes [47]. RAB27A, like MYO5A, localizes to melanosomes and is mostly concentrated in melanosome-rich dendritic tips of wild-type melanocytes. Rab27a-deﬁcient ashen melanocytes exhibit normal dendritic morphology and melanosome biogenesis but an abnormal accumulation of end-stage melanosomes in the cell center. Ménasché et al. were the ﬁrst to demonstrate that RAB27A mutations found in GS patients were linked to GS2 [48], which accounts for most cases of GS to date [26]. Re-expression of RAB27A in GS melanocytes (or ashen melanocytes) restores the proper distribution of melanosomes to the dendritic tips, stressing its importance in melanosome trafﬁcking. Coimmunoprecipitation studies with antibodies against Rab27a proved an association between Rab27a and Myo5a. This association was found to be indirect, and Mlph was later identiﬁed as the linker protein connecting both Rab27a and Myo5a (see Section 10.3.2.3). As mentioned above, the second type of GS (GS2, OMIM #607624) is caused by mutations in RAB27A, which is also located on chromosome 15q21. GS2 patients suffer from pigmentary dilution of skin and hair, and also develop immunodeﬁciency due to impaired lytic granule exocytosis in cytotoxic T lymphocytes. This can lead to episodes of a life-threatening uncontrolled T lymphocyte and macrophage activation syndrome also known as hemophagocytic syndrome or hemophagocytic lymphohistiocytosis [26]. Apart from its role in melanosome transport, the RAB27A protein is thus also involved in docking and release of lytic granules in cytotoxic T lymphocytes. 10.3.2.3 MLPH Matesic et al. identiﬁed melanophilin (Mlph) as the mutated gene in leaden (ln) mice [22]. Since the phenotype of the leaden mouse is similar to the ashen and dilute mouse, Mlph was considered as a potential new candidate gene that was mutated in GS. As perinuclear clustering of melanosomes was also observed in leaden melanocytes it was suggested that Mlph might function as part of a transport complex with Myo5a and Rab27a [22]. The newly identiﬁed Mlph or the synaptotagmin-like protein (Slp) lacking C2-a domains (Slac2-a) represented a novel class of the Slp family. All members of the Slp family contain an N-terminal Slp homology domain (SHD) consisting of two conserved potential α-helical regions (SHD1 and SHD2) often separated by two zinc ﬁnger motifs. In contrast to other Slp proteins, the Slac2 family (Slac2-a/ Mlph, Slac2-b, and Slac2-c/Myrip) lacks C-terminal tandem C2 domains (termed C2A and C2B). Instead, MLPH contains two unique coiled-coil domains (Coil 1 and Coil 2) at its C-terminal end [49]. Biochemical and cell biological analyses have now revealed that MLPH is the speciﬁc linker protein between MYO5A and RAB27A (for more details, see below).

303

304

10 Transport and Distribution of Melanosomes

In 2003, the sole mutation in the human MLPH gene (located on 2q37), giving rise to GS type 3 (GS3, OMIM #609227), was documented [50]. Interestingly, phenotypic expression of GS3 is restricted to the characteristic hypopigmentation of this syndrome. Loss of function of MLPH does not cause neurological nor immunological defects. GS3-associated hypomelanosis is indistinguishable from that described in GS1 and GS2. A second Rab27a effector was also identiﬁed in melanocytes: Slp2-a. Slp2-a contains two C2 domains and silencing experiments demonstrated that the C2A domain binds phosphatidylserine, thereby attaching melanosomes to the plasma membrane [51]. Both Rab27a effectors thus act successively in melanosome transfer: Mlph in controlling transfer from microtubules to actin ﬁlaments as well as actin-based melanosome transport and Slp2-a in anchoring of melanosomes to the plasma membrane (see also Figure 10.2). This sequential action is accompanied by differences in Rab27a binding afﬁnities, with low-afﬁnity binding of Mlph and high-afﬁnity binding of Slp2-a [52]. 10.3.2.4 RAB27A–MLPH–MYO5A Tripartite Protein Complex In the past, we have reported that human MYO5A undergoes tissue-speciﬁc alternative splicing located at its medial tail region leading to an alternate usage of three exons designated as B, D, and F [41]. Of the six known human isoforms (ABCDEF, ABCEF, ACDEF, ABCDE, ABCE, and ACE), three contain exon F. These exon F-containing isoforms are most abundantly expressed in melanocytes and were shown to colocalize with melanosomes [41, 53, 54]. Molecular studies (use of dominant-negative constructs, rescue experiments, yeast two-hybrid screenings) revealed that both the C-terminal globular tail domain and the exon F sequence are essentially required for Myo5a to colocalize with and inﬂuence the position of melanosomes in melanocytes [53, 54]. It became clear that Rab27a and Myo5a exon F-containing isoforms function as a receptor complex involving a speciﬁc linker protein. Yeast two-hybrid screenings containing exon F and globular tail constructs revealed speciﬁc interactions with a Rab effector protein termed Mlph [54, 55]. Other groups conﬁrmed these observations by performing colocalization studies in leaden or ashen melanocytes [25, 56] or in vitro binding assays [57]. The region of Mlph responsible for binding of both Rab27a and Myo5a was mapped to its N- and C-terminus, respectively [49, 55, 57]. Based on the results summarized above, the recruitment of Myo5a to melanosomes occurs as follows: activated Rab27a ﬁrst binds to the surface of the melanosomes and then recruits Mlph, which subsequently recruits the Myo5a exon F-containing isoforms (Figure 10.4a). It is now generally accepted that the Rab27a–Mlph– Myo5a tripartite complex is essential for the transportation of melanosomes from the microtubules to the actin ﬁlaments and for the retention of the melanosomes on the sublemmal actin cytoskeleton [54, 55, 57, 58]. Moreover, in vitro reconstitution of the Myo5a receptor complex by use of dominant active Rab27a and Mlph demonstrated that these proteins are not only necessary, but also sufﬁcient, to form a transport complex that moves processively on actin [59]. Loss of one of the components of the tripartite complex results in GS, biologically

10.3 Intracellular Melanosome Transport a)

b)

Figure 10.4 Functional role of the Rab27a– Mlph–Myo5a tripartite complex in intracellular melanosome transport. (a) In melanocytes, activated Rab27a is present on mature melanosomes that move to the cell periphery on microtubules via the motor protein kinesin. Once arrived at the cell periphery, Mlph is directly assembled via its SHD1 domain (part of the Rab27a-binding domain, R27BD) to the switch II region of Rab27a. Finally, Myo5a is recruited to the Rab27a–Mlph complex through a direct interaction of its globular tail and exon F sequence with two distinct regions located in middle domain of Mlph (MBD). The microtubule end-binding protein (EB-1) physically interacts with the ABD of Mlph, but its exact function in melanosome transport still needs to be elucidated.

Eventually, a stable Rab27a–Mlph–Myo5a tripartite protein complex is formed that captures the melanosomes in the actin-rich dendritic tips, a necessary step before transfer of melanosomes to the surrounding keratinocytes. Loss of function of the tripartite complex due to mutations in MYO5A, RAB27A, or MLPH/SLAC2-A leads respectively to GS types 1 (GS1), 2 (GS2), or 3 (GS3). (b) Phase-contrast microscopy images of a normal melanocyte (a) compared to a GS-derived melanocyte (b) where the abnormal perinuclear accumulation of melanosomes is depicted. Bars: 25 μm. ZnF, zinc ﬁnger; GTBD, globular tail-binding domain; EFBD, exon F-binding domain; C, coil; 1–590, the amino acid sequence of mouse MLPH. (Reproduced from [26].)

characterized by a perinuclear accumulation of melanosomes in the melanocytes (Figure 10.4b). Several binding domains (and critical residues) of Mlph have been identiﬁed as being essential for the correct binding of Rab27a and proper recruitment of Myo5a. The N-terminal part of Mlph, containing SHD1, SHD2, and an intervening zinc

305

306

10 Transport and Distribution of Melanosomes

ﬁnger domain, is responsible for interaction with Rab27a and is termed R27BD (RAB27A-binding domain). Within this region, SHD1 binds directly with the switch II region of the GTP-bound active form of Rab27a present on melanosomes [60, 61]. The globular tail and exon F sequence of Myo5a bind two distinct domains of Mlph; Myo5a-GT binds to a newly characterized region adjacent to SHD2 (mouse amino acids 147–240, termed GTBD). In comparison, Myo5a-exon F has been proven to bind to the middle region of Mlph (mouse amino acids 320–406, termed EFBD) (see also Figure 10.4a). The interaction between Mlph and Myo5aGT seems to be less stable and much weaker compared to the interaction between Mlph and Myo5a-exon F. Interestingly, studies with dilute missense mutations in Myo5a-GT impaired only the former interaction and not the latter, indicating that the Mlph–Myo5a-GT is physiologically relevant. As a consequence, both domains are essential and probably function in a synergistic manner during melanosome transport in melanocytes [62]. Hume et al. extended the results shown above by demonstrating that melanosomal Rab27a-GTP recruits Mlph to the melanosomes via interaction with both SHD1 and SHD2 [63]. In addition, the authors identiﬁed a coiled-coil structure in the C-terminal end (mouse amino acids 440–483), termed the actin-binding domain (ABD) of Mlph which is essential for the recruitment of Myo5a to the melanosomes by increasing the interaction of the Myo5a-binding domain (MBD) of Mlph with Myo5a. Additional binding partners, apart from the ones of the tripartite complex, were also identiﬁed for Mlph. The ABD of Mlph interacts with actin and with EB-1, the microtubule plus-end tracking protein (Figure 10.4a). It is suggested that interaction with actin might enhance the Myo5a motor activity [64]. The binding of EB-1 is proposed to allow transport of Mlph to the peripheral dendrite tips on the plusend tips of growing microtubules, where it would reside in complex with actinassociated Myo5a until a suitable Rab27a-associated melanosome could be captured [65]. The role of actin and EB-1 binding by Mlph in melanosome transport is disputed by recent research. Expression of Mlph mutants that lack the ABD in leaden melanocytes are able to rescue peripheral melanosome transport just like wild-type Mlph, and depletion of EB-1 using small interfering RNA does not affect melanosome transport [63, 66]. Further research will be necessary to determine if these interactions play functionally redundant roles or if they are involved in other processes (e.g., melanosome transfer). 10.3.2.5 RAB27A as a New MITF Target Gene In vitro studies in B16 mouse melanoma cells demonstrated that the α-melanocyte stimulating hormone, by means of activating the cAMP pathway, rapidly induces an increase in the interaction of Mlph with actin, resulting in a fast accumulation of melanosomes around the actin-rich region of the dendrite extremities. Additionally, cAMP stimulates the expression of Rab27a that could facilitate the interaction of melanosomes with cortical actin [67]. In melanocytes the effects of cAMP on melanin synthesis are mediated by microphtalmia-associated transcription factor (MITF) that plays an essential role in survival, migration, proliferation and differentiation of melanocytes during development. MITF controls the expression of

10.4 Melanosome Motility in RPE: The Rab27a–Myrip–Myo7a Tripartite Complex

genes essential for melanin synthesis (such as TYR (tyrosinase), TYRP1 (tyrosinaserelated protein 1), and DCT (dopachrome tautomerase; also known as TYRP2)) and for the maturation of melanosomes (MART-1, PMEL17 and GPR143 (OA1)) [68]. Interestingly, one study also demonstrated that Mitf contributes to melanosome distribution and dendrite formation of frog melanophores [69]. Based on the knowledge described above, Ballotti et al. decided to examine the possible role of MITF in melanosome transport. Silencing of MITF in human and mouse melanoma cells induced perinuclear aggregation of melanosomes, including a relocalization of RAB27A, MLPH, and MYO5A to the cell body caused by a dramatic decrease in RAB27A expression. Functional analysis of the RAB27A promoter revealed that MITF directly binds to two E-boxes in the proximal region of the RAB27A promoter, thereby stimulating the expression of RAB27A and facilitating its interaction with MLPH [70]. In this manner, loss of MITF inhibits the expression of RAB27A (and blocks the effect of cAMP on RAB27A), which consequently results in the impairment of the RAB27A–MLPH–MYO5A tripartite complex, thus explaining the abnormal melanosome distribution in MITF-deﬁcient melanoma cells. The identiﬁcation of RAB27A as a new MITF target gene links MITF to actin-dependent melanosome transport, an important step in melanocyte differentiation.

10.4 Melanosome Motility in RPE: The Rab27a–Myrip–Myo7a Tripartite Complex

In mammals, melanosomes are not solely found in epidermal melanocytes, but are also present in choroidal melanocytes of the eye and RPE cells. In the latter cell type, melanosome transport into apical processes, which surround the photoreceptor outer segments, is regulated by incident light and/or circardian rhythms [3, 29]. The molecular mechanisms involved in melanosome distribution have been examined in detail in mammalian RPE cells. Liu et al. demonstrated that RPE cells from shaker-1 mice, which carry a mutation in the gene encoding Myo7a, exhibit a perinuclear distribution of their melanosomes [71]. They are excluded from the apical processes indicating that Myo7a is required for movement. Further, Myo7a was detected on retinal melanosomes by immunoelectron microscopy [28, 71]. A human counterpart for Shaker-1 mice is found in Usher syndrome 1B and the role of Myo7a in human and mouse RPE cells is accepted as comparable [72]. Rab27a is also present on RPE melanosomes and the phenotype of ashen cells is similar to that of shaker-1 cells, with perinuclear aggregation of melanosomes [73, 74]. A Mlph analog, Myrip (also know as Slac2-c), was detected as a linker protein between Rab27a and Myo7a [28, 75]. In vitro studies revealed that Myrip acts as a multifunctional myosin-activating protein able to interact and activate both Myo7a and Myo5a, through different binding domains [76]. Conclusive evidence for the function of a Rab27a–Myrip–Myo7a tripartite complex in the transport of RPE melanosomes, similar to the function of Rab27a–Mlph–Myo5a in the

307

308

10 Transport and Distribution of Melanosomes

transport of epidermal melanosomes, came from RNA interference studies. Loss of any of the three components resulted in a redistribution of melanosomes from the actin-rich apical region and processes to the microtubule-rich cell body, the same phenotype as observed in ashen and shaker-1 cells [77]. Interestingly, nocodazole treatment (to disrupt microtubules) of RPE cells leads to an almost complete loss of melanosome movement suggesting that, as in epidermal melanocytes, melanosome transport also requires microtubular motors such as kinesins and dyneins [77]. See Figure 10.5.

Melanosome transport in mouse RPE cells. Melanosomes display fast, bidirectional, microtubule-dependent long-range movements in the cell body driven by kinesin/dynein motor proteins. Rab27a, which is targeted to the melano-

Figure 10.5

some membrane, interacts with its effector, Myrip, which in turn binds to Myo7a. Myo7a then enables the retention and/or local movement of the melanosomes along the actin ﬁlaments of the RPE apical processes.

10.5 Melanosome Transfer

10.5 Melanosome Transfer

Melanosomes need to be delivered to surrounding skin cells in order to fulﬁll their function – protecting genetic material of skin cells against UV damage. This process of delivery of melanosomes from melanocytes to surrounding skin cells (i.e., keratinocytes) is called melanosome transfer. In contrast to the extensive knowledge on the transport of melanosomes inside the melanocyte, relatively little is known about the processes and molecules involved in their transfer. Different hypotheses have been postulated. They are (i) (cyto)phagocytosis, (ii) the release of melanosomes by the melanocytes (exocytosis) and the subsequent endocytosis (or phagocytosis) of naked melanin by the keratinocytes, (ii) the active transfer of melanosome-containing vesicles by the melanocyte into the keratinocyte, and (iv) fusion of plasma membranes with tunnel formation [78]. Transfer of melanin in the hair follicle from mature melanocytes residing at the hair bulb to cortical and medullary keratinocytes is presumed to involve the same mechanisms as in the epidermal melanin unit [79]. 10.5.1 Modes of Transfer 10.5.1.1 Cytophagocytosis Phagocytosis is the cellular internalization of particles with a diameter of more than 0.5 μm. The process is receptor-mediated and activation of phagocytic receptors, such as the Fcγ receptors, leads to local reorganization of the actin cytoskeleton. Phagocytosis is mainly associated with neutrophils, macrophages, and monocytes – all “professional” phagocytes functioning to eliminate infectious agents, apoptotic cells, and cellular debris. Phagocytic activity of keratinocytes has been shown previously by use of different approaches [80–82] Cytophagocytosis stands for phagocytosis of a viable cell or an intact part of a viable cell. The cytophagocytosis hypothesis of melanin transfer describes phagocytosis of an intact melanocytic cell part (i.e., the tip of a dendrite) by the keratinocyte. At the ﬁrst stage, the melanocyte extends its dendrite towards and makes contact with a surrounding keratinocyte. The keratinocyte reacts with extensive membrane rufﬂing and engulﬁng of the dendrite tip with villus-like cytoplasmic projections. In the second stage, the dendrite tip is being squeezed and pinched off, resulting in the formation of a cytoplasmic poach ﬁlled with melanosomes. In the third stage, a phagolysosome is formed by fusion of lysosomes, degradation of the melanocyte membranes and cytoplasmic constituents takes place and, meanwhile, the phagolysosome is transported to the supranuclear region. In the fourth and last stage, the phagolysosome disintegrates into smaller vesicles containing a single melanin granule or aggregates of melanin granules, which are then dispersed over the cytoplasm (Figure 10.6a) [78]. The hypothesis of cytophagocytosis has basically been supported by electron microscopy and time-lapse video microscopy studies [78].

309

310

10 Transport and Distribution of Melanosomes a)

b)

c)

Different modes of melanosome transfer. (a) Cytophagocytosis: a melanocyte (MC) dendrite is pinched off and phagocytosed, leading to a phagolysosome from which melanosomes disperse throughout the cytoplasm. (b) Exocytosis: melanin is externalized by fusion of the melanosomal membrane with the plasma membrane and is then taken up by endocytosis or phagocy-

Figure 10.6

tosis. (Adapted from [78].) (c) Filopodialphagocytosis model: after formation and elongation, melanocyte ﬁlopodia (containing melanosomes) adhere to and are inserted into the keratinocyte (KC) plasma membrane. The melanocyte ﬁlopodia with melanosomes are ﬁnally taken up by the keratinocyte through phagocytosis. (Adapted from [4].)

10.5.1.2 Exocytosis All eukaryotic cells undergo constitutive exocytosis. However, specialized secretory cells, such as neurons, and cells of the endocrine, exocrine, and immune system show regulated exocytosis – a process in which the membranes of cytoplasmic organelles fuse with the plasma membrane in response to stimulation. The process functions to secrete products that are segregated in the organelle lumen to the extracellular space [83]. Figure 10.6(b) depicts the exocytosis process for melanin transfer: fusion of the melanosomal membrane with the melanocyte plasma membrane results in extracellular melanin, which is subsequently endocytosed or phagocytosed by surrounding keratinocytes. Evidence suggesting that (regulated) exocytosis is involved in the transfer process was derived from electron microscopic analyses of human skin and hair follicles, demonstrating naked melanin (i.e., not surrounded with a membrane) in the intercellular space and the enfolding of these pigment granules by kerati-

10.5 Melanosome Transfer

nocyte pseudopods or clathrin-coated pits [84]. Further, in vitro studies demonstrated that melanocytes discharged melanin in the extracellular space, especially when stimulated with α-melanocyte-stimulating hormone or UV [81]. Moreover, SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) proteins and Rab GTPases, involved in exocytosis in other cell types, were shown to be expressed by melanocytes. SNARE proteins are implicated in almost all intracellular membrane trafﬁcking events [85]. The synaptic proteins syntaxin (STX), SNAP25 (25-kDa synaptosome-associated protein), and VAMP (vesicle-associated membrane protein, also called synaptobrevin) were the ﬁrst SNAREs to be discovered. These three conserved SNARE families act late in the events of membrane fusion. They associate into core complexes, and usually SNAP25 and syntaxin on the plasma membrane bind VAMP on the vesicle membrane. Different SNAREs were identiﬁed in melanosome-enriched fractions: SNAP23, SNAP25, VAMP2, STX4, and STX6 [86, 87]. Immunoprecipitation shows association of VAMP2 and SNAP23, but not STX4. Possibly, VAMP2 on melanosomes interacts with SNAP23 and an as yet to be identiﬁed syntaxin on the melanocyte plasma membrane to achieve fusion. Other proteins that play a role in membrane fusion, particularly in tethering and docking of membranes before actual fusion, are Rab GTPases. The Rab3 proteins consisting of Rab3a–d are the central Rabs in regulated exocytosis. Rab3a is expressed in melanocytes and downregulation of its expression is effected by UV irradiation [86]. Interestingly, downregulation of Rab3 in other cell types has proved to stimulate regulated exocytosis [88, 89]. 10.5.1.3 Filopodial-Phagocytosis Model Filopodia are narrow (200- to 300-nm diameter) highly motile tubular membrane extensions that contain long actin ﬁlaments organized as bundles, with plus-ends/ barbed ends (fast-growing ends) in the direction of protrusions. Filopodia seem to be used by many cell types as a sensing organelle to explore the extracellular matrix and the surface of other cells, to identify appropriate targets for adhesion, and then generate guidance cues and traction forces to move the cell body. Several studies revealed their involvement in the transport of different organelles such as membrane vesicles [90–93]. Filopodia extent from the dendrite tips and cell body of melanocytes, adhere to the surface of neighboring keratinocytes, and allow transport of melanosomes towards the keratinocyte membrane. In some instances, transfer was seen via these protrusions, suggesting the formation of a tunnel between melanocyte and keratinocyte cytoplasms [94, 95]. Deﬁnitive proof of membrane fusion could not be given, though. The role of ﬁlopodia in melanosome transfer was further explored in great detail by Singh et al. Scanning electron microscopy studies of epidermal melanocyte–keratinocyte cocultures revealed numerous intraluminal ovoid bodies present in ﬁlopodia extending from melanocytes to abut adjacent keratinocytes. These bodies were shown to be gp100-positive and were thereby identiﬁed as melanosomes [4]. Time-lapse video micrography conﬁrmed their transfer from melanocytes to keratinocytes. Interestingly, transfer was inhibited

311

312

10 Transport and Distribution of Melanosomes

when silencing a regulator of ﬁlopodium formation, MyoX [96, 97], in melanocytes, and when blocking ﬁlopodium formation by addition of low-dose cytochalasin B. As MyoX is also an effector of phagocytosis [98], Singh et al. decided to examine its role in the interaction of the melanocyte ﬁlopodia with the keratinocyte plasma membrane – an interaction that is crucial for the successful transfer of melanosomes. Knockdown of MyoX in keratinocytes resulted in an almost complete inhibition of melanosome uptake by keratinocytes. Based on these ﬁndings the authors suggested a dual role for MyoX in melanosome transfer: formation of ﬁlopodia in melanocytes and subsequent phagocytosis of ﬁlopodia by keratinocytes. This new model of melanosome transfer between human skin cells was termed the “ﬁlopodial-phagocytosis” model (Figure 10.6c) [4]. The same authors reported the unexpected ﬁnding that keratinocyte ﬁlopodia are also conduits for melanin between adjacent keratinocytes. At the moment one can only speculate about the role(s) of this homotypic melanin transfer. It might, for example, facilitate melanin degradation – a process largely uncharacterized. Further research will be necessary to sustain such or other hypotheses. 10.5.2 Molecular Players 10.5.2.1 PAR-2 and KGF Protease activated receptor-2 (PAR-2) belongs to a family of transmembrane G-protein-coupled receptors (PAR-1 to -4) that are proteolytically activated by serine proteases. These enzymes (including trypsin or mast cell tryptase) cleave the extracellular amino terminal domain of PAR-2. The newly formed N-termini are tethered ligands, they undergo a conformational change and bind the receptors leading to activation. The use of synthetic peptides, corresponding to the sequence of the N-terminus of the cleaved receptor, also enables activation of PAR-2 independent of receptor cleavage. A role for PAR-2 in melanin transfer has been established by the group of Seiberg [99]. They showed that stimulation of this receptor, which is expressed in keratinocytes [100], but not in melanocytes [101], enhances the phagocytosis rate of keratinocytes and leads to increased melanin transfer [101–103]. Melanocyte–keratinocyte contact is a prerequisite for this function [101]. UV irradiation induces PAR-2 and, conversely, blocking of the PAR-2 receptor inhibits UV-induced pigmentation [104]. Further, PAR-2 expression and induction by UV seem to depend on skin type, with a higher expression and more pronounced induction in dark-skinned individuals [105]. It has been shown in vitro that activation of PAR-2 leads to serine protease secretion by keratinocytes, creating a positive feedback loop. UV-B induces a similar effect [104]. However, melanin transfer cannot be completely inhibited by treatment with serine protease inhibitors [102]. This suggests that PAR-2 is probably not the only molecular player involved in phagocytosis by keratinocytes. In fact, the keratinocyte growth factor receptor (KGFR) has been allocated a similar role [106]. Activation of KGFR enhances keratinocyte phagocytosis of latex beads and addition of KGF to cocultures induces transfer of tyrosinase-positive granules.

10.6 Fate of Melanin in the Keratinocyte

Apart from phagocytosis, PAR-2 affects skin pigmentation by stimulation of melanocyte dendricity. Prostaglandins E2 and F2α are released by keratinocyte upon stimulation. They bind the surface of melanocytes, thereby inducing dendrite formation [107]. 10.5.2.2 Adhesion Molecules: Cadherins and Lectins Cell–cell contact between melanocytes and keratinocytes is a prerequisite for melanosome transfer. Different adhesion molecules are involved at the melanocyte– keratinocyte contact site. Cadherins are a family of glycoproteins that function in promoting Ca2+-dependent cell–cell adhesion and serve as the transmembrane components of cell–cell adherent junctions. E-cadherin and P-cadherin are expressed in human melanocytes, and both mediate melanocyte adhesion to keratinocytes. P-cadherin seems to play a minor role as opposed to E-cadherin, which is the major mediator of melanocyte–keratinocyte adhesion [108]. A role for E-cadherin in pigment transfer has been suggested based on ﬁndings in Darier’s disease. This acantholytic disorder, caused by a mutation in the (sarco)endoplasmic reticulum pump SERCA2, is sometimes associated with confetti-like hypopigmented macules in dark-skinned individuals. The ultrastructure of these hypopigmented lesions, with basal and suprabasal keratinocytes being “empty” despite being surrounded by melanosome-ﬁlled dendrites, suggests a defect in melanosome transfer. Considering the importance of E-cadherin in melanocyte– keratinocyte adhesion and the fact that it is dissociated in acantholytic lesions of Darier’s disease points to a possible involvement of E-cadherin in the transfer process [109]. Lectins are adhesion receptors that bind sugar residues present on the surface of delimiting membranes. When added to keratinocyte–melanocyte cultures, lectins and neoglycoproteins inhibit melanin transfer, as shown by ﬂow cytometry and electron microscopy [110]. The inhibition is reversible and can be enhanced by addition of niacinamide [111]. Lectins that bind galactose residues are more effective than lectins binding mannose residues, while α-l-fucose receptors display a variable effect. Cerdan et al. studied the role of these molecules in binding of melanin-containing vesicles shed by melanoma cells to keratinocytes [112]. Binding is inhibited by neoglycoproteins, showing a role for α-l-fucose-speciﬁc lectins of keratinocytes, on the one hand, and for 6-phospho-β-d-galactose-speciﬁc lectins of melanocyte-derived vesicles, on the other hand.

10.6 Fate of Melanin in the Keratinocyte

Once transferred, melanin granules distribute differently according to the skin phototype. In light skin, melanin granules cluster in membrane-bound organelles that border on the nucleus, while in dark-skinned individuals melanin granules are distributed throughout the cell body [113]. Light skin predominantly contains smaller melanosomes than dark skin (around 0.5 versus 0.8 μm in diameter) and

313

314

10 Transport and Distribution of Melanosomes

traditionally the distribution pattern was considered to be size-dependent. Small melanosomes cluster and form melanosome complexes, whereas larger ones are packed individually [114]. The size-dependency of the distribution pattern was contradicted by later studies. Analysis of cocultures composed of cells derived from different phototypes showed that the origin of the recipient keratinocyte is the decisive factor [114]. The signiﬁcant role of keratinocytes in the determination of skin pigmentation became even more evident when studying human skin reconstructs composed of cells derived from different skin phototypes. Reconstructs with light skin-derived melanocytes appeared darker, produced more melanogenic cytokines as well as melanin, and contained more mature melanosomes, which had a higher individual/clustered distribution ratio, when the keratinocytes originated from dark skin versus light skin [114, 115]. Apparently, the keratinocyte inﬂuences the quality of melanosome production and it determines the distribution pattern of melanosomes after transfer. Keratinocytes can adopt a pigment recipient phenotype, leading to recruitment of melanocytes and inducing their own pigmentation. At least this was shown for epidermal keratinocytes in mouse skin. When the transcription factor Foxn1 is transgenically expressed in basal keratinocytes of mouse skin, numerous melanocytes are found in the interfollicular epidermis and melanization of this epidermis follows (whereas normally murine interfollicular epidermis is not pigmented) [116]. Keratinocytes are also crucial for the response of melanocytes to UV irradiation, and thus for the determination of facultative skin pigmentation [117]. Increased melanogenesis, for example, requires a 10-fold higher UV-B dose when melanocytes are cultured alone versus together with keratinocytes [117]. Inside keratinocytes, melanin is stored in melano-phagosomes that eventually fuse with lysosomes to form melano-phagolysosomes. Aggregation of melanophagolysosomes above and adjacent to the nucleus of suprabasal keratinocytes results in the formation of the “microparasol” or “supranuclear melanin cap,” which protects keratinocyte DNA from UV damage [118]. Supranuclear caps are also formed in cultured skin equivalents and their formation increases after UV irradiation and 3-isobutyl-1-methylxanthine treatment [119]. Transport to the cell center is accomplished by the microtubule-dependent minus-end motor protein dynein that attaches to the melano-phagolysosome via the dynactin complex [120]. Knockdown of the p150Glued subunit of dynactin, that binds dynein as well as microtubules, impairs the perinuclear targeting of ingested polystyrene microspheres in keratinocytes [121]. This underlines the importance of the dynein–dynactin complex for the formation and maintenance of the microparasol, and indicates that the transport is independent of any melanosomal component or signal, since phagocytosed microspheres behave just like melano-phagolysosomes. Data on melanin and melanosome degradation are limited. Melanosomes mainly populate the basal layers of fair-skinned as well as dark-skinned individuals and they are no longer visible in the differentiated keratinocytes from the upper epidermal layers. Only in dark skin types can some scattered melanosomes be

10.7 Conclusions

distinguished up to the stratum corneum. Hydrolytic reactions are responsible for the breakdown of the lipid and protein components of melanosomes, but melanin is a hardy, resistant structure and degradation via hydrolysis would be impossible [122]. Oxidative reactions could be responsible, for example, via NADPH oxidase, which resides on the phagosomal membrane. There is still doubt, though, on whether melanin as a chemical compound is degraded in vivo.

10.7 Conclusions

Skin pigmentation depends on the amount and type of melanin synthesized by the melanocyte, the subsequent transfer of melanosomes to the keratinocytes, and eventually the distribution of pigment in the skin. Prior to the transfer of melanosomes to keratinocytes, mature melanosomes need to be moved from the perinuclear area to the cell periphery and dendritic tips of the melanocytes. This intracellular melanosome transport is dependent on an intact cytoskeleton, and is functional in pigment cells derived from lower vertebrates and mammals. Detailed studies in both melanophores and melanocytes revealed that melanosomes undergo transport by means of a coupled system of microtubule-dependent transport to the periphery, followed by actin-dependent retention. Three different classes of motor protein were shown to be involved in this movement: kinesins, dyneins, and Myo5a. In mammals, melanosomes are captured below the sublemmal actin network through the formation of a Rab27a–Mlph–Myo5a tripartite protein complex. From there, melanosomes have to pass from the dendrites of the melanocytes to neighboring keratinocytes. This transfer process appears difﬁcult to unravel and the mechanisms involved remain poorly understood. The development of reliable coculture assays to study melanosome transfer, combined with high-resolution microscopy, provides growing evidence that ﬁlopodia from melanocytes are involved in melanosome transfer [4]. Moreover, ﬁlopodia were also identiﬁed as conduits for melanosome transfer between individual keratinocytes. This unexpected result might point at a much greater role for keratinocytes in the control of mammalian skin pigmentation than hitherto assumed. Interestingly, the use of long-term model systems (e.g., grafting of melanocytes and keratinocytes derived from skin of different racial/ethnic origin onto SCID mice) revealed that keratinocytes play an important role in determining the amount and type of melanin produced by melanocytes, the number of melanosomes that are transferred, and their eventually distribution pattern in keratinocytes [115]. Today, there is great interest to develop new tools in order to darken or lighten skin coloration or to treat skin pigmentary disorders in an efﬁcient and safe manner. In this context, our group is focusing on the development of RNA interference-based therapeutics in order to reduce pigmentation by silencing the tyrosinase gene, for example, or genes involved in the intracellular melanosome transport pathway. As novel insights into the mechanisms of melanosome transfer

315

316

10 Transport and Distribution of Melanosomes

become available, it will be interesting to interfere at this particular level of the pigmentary process in order to modulate pigmentation.

Acknowledgments

We would like to thank Dr Karolien Van Den Bossche for critical comments on this text and Raphael van den Eerenbeemt for drawing Figure 10.5. M.V.G. is a postdoctoral researcher of the Fund for Scientiﬁc Research-Flanders (Belgium).

References 1 Raposo, G. and Marks, M.S. (2007)

2

3

4

5

6

7

8

Melanosomes – dark organelles enlighten endosomal membrane transport. Nat. Rev. Mol. Cell. Biol., 8, 786–797. Aspengren, S., Hedberg, D., Skold, H.N., and Wallin, M. (2009) New insights into melanosome transport in vertebrate pigment cells. Int. Rev. Cell. Mol. Biol., 272, 245–302. Futter, C.E. (2006) The molecular regulation of organelle transport in mammalian retinal pigment epithelial cells. Pigment Cell Res., 19, 104–111. Singh, S.K., Kurfurst, R., Nizard, C., Schnebert, S., Perrier, E., and Tobin, D.J. (2010) Melanin transfer in human skin cells is mediated by ﬁlopodia – a model for homotypic and heterotypic lysosome-related organelle transfer. FASEB J., 24, 3756–3769. Aspengren, S., Hedberg, D., and Wallin, M. (2007) Melanophores: a model system for neuronal transport and exocytosis? J. Neurosci. Res., 85, 2591–2600. Wu, X., Rao, K., Bowers, M.B., Copeland, N.G., Jenkins, N.A., and Hammer, J.A., 3rd (2001) Rab27a enables myosin Va-dependent melanosome capture by recruiting the myosin to the organelle. J. Cell Sci., 114, 1091–1100. Daniolos, A., Lerner, A.B., and Lerner, M.R. (1990) Action of light on frog pigment cells in culture. Pigment Cell Res., 3, 38–43. Clark, T.G. and Rosenbaum, J.L. (1982) Pigment particle translocation in detergent-permeabilized melanophores

9

10

11

12

13

14

15

16

of Fundulus heteroclitus. Proc. Natl. Acad. Sci. USA, 79, 4655–4659. Grundstrom, N., Karlsson, J.O., and Andersson, R.G. (1985) The control of granule movement in ﬁsh melanophores. Acta Physiol. Scand., 125, 415–421. Nilsson, H., Rutberg, M., and Wallin, M. (1996) Localization of kinesin and cytoplasmic dynein in cultured melanophores from Atlantic cod, Gadus morhua. Cell Motil. Cytoskeleton, 33, 183–196. Rogers, S.L., Tint, I.S., Fanapour, P.C., and Gelfand, V.I. (1997) Regulated bidirectional motility of melanophore pigment granules along microtubules in vitro. Proc. Natl. Acad. Sci. USA, 94, 3720–3725. Nilsson, H. and Wallin, M. (1997) Evidence for several roles of dynein in pigment transport in melanophores. Cell Motil. Cytoskeleton, 38, 397–409. Tuma, M.C., Zill, A., Le Bot, N., Vernos, I., and Gelfand, V. (1998) Heterotrimeric kinesin II is the microtubule motor protein responsible for pigment dispersion in Xenopus melanophores. J. Cell Biol., 143, 1547–1558. Tuma, M.C. and Gelfand, V.I. (1999) Molecular mechanisms of pigment transport in melanophores. Pigment Cell Res., 12, 283–294. Schliwa, M. and Euteneuer, U. (1978) A microtuble-independent component may be involved in granule transport in pigment cells. Nature, 273, 556–558. McGuire, J., Moellmann, G., and McKeon, F. (1972) Cytochalasin B and

References

17

18

19

20

21

22

23

24

pigment granule translocation. Cytochalasin B reverses and prevents pigment granule dispersion caused by dibutyryl cyclic AMP and theophylline in Rana pipiens melanocytes. J. Cell Biol., 52, 754–758. Aspengren, S., Wielbass, L., and Wallin, M. (2006) Effects of acrylamide, latrunculin, and nocodazole on intracellular transport and cytoskeletal organization in melanophores. Cell Motil. Cytoskeleton, 63, 423–436. Schliwa, M., Weber, K., and Porter, K.R. (1981) Localization and organization of actin in melanophores. J. Cell Biol., 89, 267–275. Rogers, S.L., Karcher, R.L., Roland, J.T., Minin, A.A., Steffen, W., and Gelfand, V.I. (1999) Regulation of melanosome movement in the cell cycle by reversible association with myosin V. J. Cell Biol., 146, 1265–1276. Skold, H.N., Norstrom, E., and Wallin, M. (2002) Regulatory control of both microtubule- and actin-dependent ﬁsh melanosome movement. Pigment Cell Res., 15, 357–366. Nascimento, A.A., Roland, J.T., and Gelfand, V.I. (2003) Pigment cells: a model for the study of organelle transport. Annu. Rev. Cell Dev. Biol., 19, 469–491. Matesic, L.E., Yip, R., Reuss, A.E., Swing, D.A., O’Sullivan, T.N., Fletcher, C.F., Copeland, N.G., and Jenkins, N.A. (2001) Mutations in Mlph, encoding a member of the Rab effector family, cause the melanosome transport defects observed in leaden mice. Proc. Natl. Acad. Sci. USA, 98, 10238–10243. Mercer, J.A., Seperack, P.K., Strobel, M.C., Copeland, N.G., and Jenkins, N.A. (1991) Novel myosin heavy chain encoded by murine dilute coat colour locus. Nature, 349, 709–713. Wilson, S.M., Yip, R., Swing, D.A., O’Sullivan, T.N., Zhang, Y., Novak, E.K., Swank, R.T., Russell, L.B., Copeland, N.G., and Jenkins, N.A. (2000) A mutation in Rab27a causes the vesicle transport defects observed in ashen mice. Proc. Natl. Acad. Sci. USA, 97, 7933–7938.

25 Hume, A.N., Collinson, L.M., Hopkins,

26

27

28

29

30

31

32

33

C.R., Strom, M., Barral, D.C., Bossi, G., Grifﬁths, G.M., and Seabra, M.C. (2002) The leaden gene product is required with Rab27a to recruit myosin Va to melanosomes in melanocytes. Trafﬁc, 3, 193–202. Van Gele, M., Dynoodt, P., and Lambert, J. (2009) Griscelli syndrome: a model system to study vesicular trafﬁcking. Pigment Cell Melanoma Res., 22, 268–282. McNeil, E.L., Tacelosky, D., Basciano, P., Biallas, B., Williams, R., Damiani, P., Deacon, S., Fox, C., Stewart, B., Petruzzi, N., Osborn, C., Klinger, K., Sellers, J.R., and Smith, C.K. (2004) Actin-dependent motility of melanosomes from ﬁsh retinal pigment epithelial (RPE) cells investigated using in vitro motility assays. Cell Motil. Cytoskeleton, 58, 71–82. El-Amraoui, A., Schonn, J.S., KusselAndermann, P., Blanchard, S., Desnos, C., Henry, J.P., Wolfrum, U., Darchen, F., and Petit, C. (2002) MyRIP, a novel Rab effector, enables myosin VIIa recruitment to retinal melanosomes. EMBO Rep., 3, 463–470. Coudrier, E. (2007) Myosins in melanocytes: to move or not to move? Pigment Cell Res., 20, 153–160. Lambert, J., Vancoillie, G., and Naeyaert, J.M. (1999) Molecular motors and their role in pigmentation. Cell Mol. Biol., 45, 905–918. Westbroek, W., Lambert, J., and Naeyaert, J.M. (2001) The dilute locus and Griscelli syndrome: gateways towards a better understanding of melanosome transport. Pigment Cell Res., 14, 320–327. Hirokawa, N., Noda, Y., Tanaka, Y., and Niwa, S. (2009) Kinesin superfamily motor proteins and intracellular transport. Nat. Rev. Mol. Cell Biol., 10, 682–696. Vancoillie, G., Lambert, J., Mulder, A., Koerten, H.K., Mommaas, A.M., Van Oostveldt, P., and Naeyaert, J.M. (2000) Cytoplasmic dynein colocalizes with melanosomes in normal human melanocytes. Br. J. Dermatol., 143, 298–306.

317

318

10 Transport and Distribution of Melanosomes 34 Vancoillie, G., Lambert, J., Mulder, A.,

35

36

37

38

39

40

41

42

Koerten, H.K., Mommaas, A.M., Van Oostveldt, P., and Naeyaert, J.M. (2000) Kinesin and kinectin can associate with the melanosomal surface and form a link with microtubules in normal human melanocytes. J. Invest. Dermatol., 114, 421–429. Vancoillie, G., Lambert, J., Haeghen, Y.V., Westbroek, W., Mulder, A., Koerten, H.K., Mommaas, A.M., Van Oostveldt, P., and Naeyaert, J.M. (2000) Colocalization of dynactin subunits P150Glued and P50 with melanosomes in normal human melanocytes. Pigment Cell Res., 13, 449–457. Gross, S.P., Welte, M.A., Block, S.M., and Wieschaus, E.F. (2002) Coordination of opposite-polarity microtubule motors. J. Cell Biol., 156, 715–724. Jordens, I., Westbroek, W., Marsman, M., Rocha, N., Mommaas, M., Huizing, M., Lambert, J., Naeyaert, J.M., and Neefjes, J. (2006) Rab7 and Rab27a control two motor protein activities involved in melanosomal transport. Pigment Cell Res., 19, 412–423. Watabe, H., Valencia, J.C., Le Pape, E., Yamaguchi, Y., Nakamura, M., Rouzaud, F., Hoashi, T., Kawa, Y., Mizoguchi, M., and Hearing, V.J. (2008) Involvement of dynein and spectrin with early melanosome transport and melanosomal protein trafﬁcking. J. Invest. Dermatol., 128, 162–174. Lakkaraju, A., Carvajal-Gonzalez, J.M., and Rodriguez-Boulan, E. (2009) It takes two to tango to the melanosome. J. Cell Biol., 187, 161–163. Trybus, K.M. (2008) Myosin V from head to tail. Cell Mol. Life Sci., 65, 1378–1389. Lambert, J., Naeyaert, J.M., Callens, T., De Paepe, A., and Messiaen, L. (1998) Human myosin V gene produces different transcripts in a cell typespeciﬁc manner. Biochem. Biophys. Res. Commun., 252, 329–333. Pastural, E., Barrat, F.J., DufourcqLagelouse, R., Certain, S., Sanal, O., Jabado, N., Seger, R., Griscelli, C., Fischer, A., and de Saint Basile, G. (1997) Griscelli disease maps to chromosome 15q21 and is associated

43

44

45

46

47

48

49

50

with mutations in the myosin-Va gene. Nat. Genet., 16, 289–292. Pastural, E., Ersoy, F., Yalman, N., Wulffraat, N., Grillo, E., Ozkinay, F., Tezcan, I., Gedikoglu, G., Philippe, N., Fischer, A., and de Saint Basile, G. (2000) Two genes are responsible for Griscelli syndrome at the same 15q21 locus. Genomics, 63, 299–306. Lambert, J., Naeyaert, J.M., De Paepe, A., Van Coster, R., Ferster, A., Song, M., and Messiaen, L. (2000) Arg–Cys substitution at codon 1246 of the human myosin Va gene is not associated with Griscelli syndrome. J. Invest. Dermatol., 114, 731–733. Corbeel, L. and Freson, K. (2008) Rab proteins and Rab-associated proteins: major actors in the mechanism of protein-trafﬁcking disorders. Eur. J. Pediatr., 167, 723–729. Figueiredo, A.C., Wasmeier, C., Tarafder, A.K., Ramalho, J.S., Baron, R.A., and Seabra, M.C. (2008) Rab3GEP is the non-redundant guanine nucleotide exchange factor for Rab27a in melanocytes. J. Biol. Chem., 283, 23209–23216. Fukuda, M. (2005) Versatile role of Rab27 in membrane trafﬁcking: focus on the Rab27 effector families. J. Biochem., 137, 9–16. Ménasché, G., Pastural, E., Feldmann, J., Certain, S., Ersoy, F., Dupuis, S., Wulffraat, N., Bianchi, D., Fischer, A., Le Deist, F., and de Saint Basile, G. (2000) Mutations in RAB27A cause Griscelli syndrome associated with haemophagocytic syndrome. Nat. Genet., 25, 173–176. Nagashima, K., Torii, S., Yi, Z., Igarashi, M., Okamoto, K., Takeuchi, T., and Izumi, T. (2002) Melanophilin directly links Rab27a and myosin Va through its distinct coiled-coil regions. FEBS Lett., 517, 233–238. Ménasché, G., Ho, C.H., Sanal, O., Feldmann, J., Tezcan, I., Ersoy, F., Houdusse, A., Fischer, A., and de Saint Basile, G. (2003) Griscelli syndrome restricted to hypopigmentation results from a melanophilin defect (GS3) or a MYO5A F-exon deletion (GS1). J. Clin. Invest., 112, 450–456.

References 51 Kuroda, T.S. and Fukuda, M. (2004)

52

53

54

55

56

57

58

59

60

Rab27A-binding protein Slp2-a is required for peripheral melanosome distribution and elongated cell shape in melanocytes. Nat. Cell Biol., 6, 1195–1203. Fukuda, M. (2006) Distinct Rab27A binding afﬁnities of Slp2-a and Slac2-a/ melanophilin: hierarchy of Rab27A effectors. Biochem. Biophys. Res. Commun., 343, 666–674. Wu, X., Wang, F., Rao, K., Sellers, J.R., and Hammer, J.A., 3rd (2002) Rab27a is an essential component of melanosome receptor for myosin Va. Mol. Biol. Cell, 13, 1735–1749. Westbroek, W., Lambert, J., Bahadoran, P., Busca, R., Herteleer, M.C., Smit, N., Mommaas, M., Ballotti, R., and Naeyaert, J.M. (2003) Interactions of human myosin Va isoforms, endogenously expressed in human melanocytes, are tightly regulated by the tail domain. J. Invest. Dermatol., 120, 465–475. Wu, X.S., Rao, K., Zhang, H., Wang, F., Sellers, J.R., Matesic, L.E., Copeland, N.G., Jenkins, N.A., and Hammer, J.A., 3rd (2002) Identiﬁcation of an organelle receptor for myosin-Va. Nat. Cell Biol., 4, 271–278. Provance, D.W., James, T.L., and Mercer, J.A. (2002) Melanophilin, the product of the leaden locus, is required for targeting of myosin-Va to melanosomes. Trafﬁc, 3, 124–132. Fukuda, M., Kuroda, T.S., and Mikoshiba, K. (2002) Slac2-a/ melanophilin, the missing link between Rab27 and myosin Va: implications of a tripartite protein complex for melanosome transport. J. Biol. Chem., 277, 12432–12436. Goud, B. (2002) How Rab proteins link motors to membranes. Nat. Cell Biol., 4, E77–E78. Wu, X., Sakamoto, T., Zhang, F., Sellers, J.R., and Hammer, J.A., 3rd (2006) In vitro reconstitution of a transport complex containing Rab27a, melanophilin and myosin Va. FEBS Lett., 580, 5863–5868. Fukuda, M. (2002) Synaptotagmin-like protein (Slp) homology domain 1 of

61

62

63

64

65

66

67

68

69

Slac2-a/melanophilin is a critical determinant of GTP-dependent speciﬁc binding to Rab27A. J. Biol. Chem., 277, 40118–40124. Strom, M., Hume, A.N., Tarafder, A.K., Barkagianni, E., and Seabra, M.C. (2002) A family of Rab27-binding proteins. Melanophilin links Rab27a and myosin Va function in melanosome transport. J. Biol. Chem., 277, 25423–25430. Fukuda, M. and Kuroda, T.S. (2004) Missense mutations in the globular tail of myosin-Va in dilute mice partially impair binding of Slac2-a/melanophilin. J. Cell Sci., 117, 583–591. Hume, A.N., Tarafder, A.K., Ramalho, J.S., Sviderskaya, E.V., and Seabra, M.C. (2006) A coiled-coil domain of melanophilin is essential for myosin Va recruitment and melanosome transport in melanocytes. Mol. Biol. Cell, 17, 4720–4735. Kuroda, T.S., Ariga, H., and Fukuda, M. (2003) The actin-binding domain of Slac2-a/melanophilin is required for melanosome distribution in melanocytes. Mol. Cell Biol., 23, 5245–5255. Wu, X.S., Tsan, G.L., and Hammer, J.A., 3rd (2005) Melanophilin and myosin Va track the microtubule plus end on EB1. J. Cell Biol., 171, 201–207. Hume, A.N., Ushakov, D.S., Tarafder, A.K., Ferenczi, M.A., and Seabra, M.C. (2007) Rab27a and MyoVa are the primary Mlph interactors regulating melanosome transport in melanocytes. J. Cell Sci., 120, 3111–3122. Passeron, T., Bahadoran, P., Bertolotto, C., Chiaverini, C., Busca, R., Valony, G., Bille, K., Ortonne, J.P., and Ballotti, R. (2004) Cyclic AMP promotes a peripheral distribution of melanosomes and stimulates melanophilin/Slac2-a and actin association. FASEB J., 18, 989–991. Vachtenheim, J. and Borovansky, J. (2010) “Transcription physiology” of pigment formation in melanocytes: central role of MITF. Exp. Dermatol., 19, 617–627. Kawasaki, A., Kumasaka, M., Satoh, A., Suzuki, M., Tamura, K., Goto, T., Asashima, M., and Yamamoto, H.

319

320

10 Transport and Distribution of Melanosomes

70

71

72

73

74

75

76

77

(2008) Mitf contributes to melanosome distribution and melanophore dendricity. Pigment Cell Melanoma Res., 21, 56–62. Chiaverini, C., Beuret, L., Flori, E., Busca, R., Abbe, P., Bille, K., Bahadoran, P., Ortonne, J.P., Bertolotto, C., and Ballotti, R. (2008) Microphthalmiaassociated transcription factor regulates RAB27A gene expression and controls melanosome transport. J. Biol. Chem., 283, 12635–12642. Liu, X., Ondek, B., and Williams, D.S. (1998) Mutant myosin VIIa causes defective melanosome distribution in the RPE of shaker-1 mice. Nat. Genet., 19, 117–118. Gibbs, D., Diemer, T., Khanobdee, K., Hu, J., Bok, D., and Williams, D.S. (2010) Function of MYO7A in the human RPE and the validity of shaker1 mice as a model for Usher syndrome 1B. Invest. Ophthalmol. Vis. Sci., 51, 1130–1135. Futter, C.E., Ramalho, J.S., Jaissle, G.B., Seeliger, M.W., and Seabra, M.C. (2004) The role of Rab27a in the regulation of melanosome distribution within retinal pigment epithelial cells. Mol. Biol. Cell, 15, 2264–2275. Gibbs, D., Azarian, S.M., Lillo, C., Kitamoto, J., Klomp, A.E., Steel, K.P., Libby, R.T., and Williams, D.S. (2004) Role of myosin VIIa and Rab27a in the motility and localization of RPE melanosomes. J. Cell Sci., 117, 6473–6483. Fukuda, M. and Kuroda, T.S. (2002) Slac2-c (synaptotagmin-like protein homologue lacking C2 domains-c), a novel linker protein that interacts with Rab27, myosin Va/VIIa, and actin. J. Biol. Chem., 277, 43096–43103. Ramalho, J.S., Lopes, V.S., Tarafder, A.K., Seabra, M.C., and Hume, A.N. (2009) Myrip uses distinct domains in the cellular activation of myosin VA and myosin VIIA in melanosome transport. Pigment Cell Melanoma Res., 22, 461–473. Lopes, V.S., Ramalho, J.S., Owen, D.M., Karl, M.O., Strauss, O., Futter, C.E., and Seabra, M.C. (2007) The ternary Rab27a–Myrip–myosin VIIa complex

78

79

80

81

82

83

84

85

86

87

regulates melanosome motility in the retinal pigment epithelium. Trafﬁc, 8, 486–499. Van Den Bossche, K., Naeyaert, J.M., and Lambert, J. (2006) The quest for the mechanism of melanin transfer. Trafﬁc, 7, 769–778. Slominski, A., Wortsman, J., Plonka, P.M., Schallreuter, K.U., Paus, R., and Tobin, D.J. (2005) Hair follicle pigmentation. J. Invest. Dermatol., 124, 13–21. Wolff, K. and Konrad, K. (1972) Phagocytosis of latex beads by epidermal keratinocytes in vivo. J. Ultrastruct. Res., 39, 262–280. Virador, V.M., Muller, J., Wu, X., Abdel-Malek, Z.A., Yu, Z.X., Ferrans, V.J., Kobayashi, N., Wakamatsu, K., Ito, S., Hammer, J.A., and Hearing, V.J. (2002) Inﬂuence of alpha-melanocytestimulating hormone and ultraviolet radiation on the transfer of melanosomes to keratinocytes. FASEB J., 16, 105–107. Ando, H., Niki, Y., Yoshida, M., Ito, M., Akiyama, K., Kim, J.H., Yoon, T.J., Lee, J.H., Matsui, M.S., and Ichihashi, M. (2010) Keratinocytes in culture accumulate phagocytosed melanosomes in the perinuclear area. Pigment Cell Melanoma Res., 23, 129–133. Burgoyne, R.D. and Morgan, A. (2003) Secretory granule exocytosis. Physiol. Rev., 83, 581–632. Yamamoto, O. and Bhawan, J. (1994) Three modes of melanosome transfers in Caucasian facial skin: hypothesis based on an ultrastructural study. Pigment Cell Res., 7, 158–169. Chen, Y.A. and Scheller, R.H. (2001) SNARE-mediated membrane fusion. Nat. Rev. Mol. Cell. Biol., 2, 98–106. Scott, G. and Zhao, Q. (2001) Rab3a and SNARE proteins: potential regulators of melanosome movement. J. Invest. Dermatol., 116, 296–304. Wade, N., Bryant, N.J., Connolly, L.M., Simpson, R.J., Luzio, J.P., Piper, R.C., and James, D.E. (2001) Syntaxin 7 complexes with mouse Vps10p tail interactor 1b, syntaxin 6, vesicleassociated membrane protein (VAMP)8,

References

88

89

90

91

92

93

94

95

96

97

and VAMP7 in b16 melanoma cells. J. Biol. Chem., 276, 19820–19827. Schluter, O.M., Khvotchev, M., Jahn, R., and Sudhof, T.C. (2002) Localization versus function of Rab3 proteins. Evidence for a common regulatory role in controlling fusion. J. Biol. Chem., 277, 40919–40929. Martelli, A.M., Baldini, G., Tabellini, G., Koticha, D., Bareggi, R., and Baldini, G. (2000) Rab3A and Rab3D control the total granule number and the fraction of granules docked at the plasma membrane in PC12 cells. Trafﬁc, 1, 976–986. Rustom, A., Saffrich, R., Markovic, I., Walther, P., and Gerdes, H.H. (2004) Nanotubular highways for intercellular organelle transport. Science, 303, 1007–1010. Onfelt, B., Nedvetzki, S., Yanagi, K., and Davis, D.M. (2004) Cutting edge: Membrane nanotubes connect immune cells. J. Immunol., 173, 1511–1513. Vidulescu, C., Clejan, S., and O’Connor, K.C. (2004) Vesicle trafﬁc through intercellular bridges in DU 145 human prostate cancer cells. J. Cell Mol. Med., 8, 388–396. Baluska, F., Hlavacka, A., Volkmann, D., and Menzel, D. (2004) Getting connected: actin-based cell-to-cell channels in plants and animals. Trends Cell Biol., 14, 404–408. Scott, G., Leopardi, S., Printup, S., and Madden, B.C. (2002) Filopodia are conduits for melanosome transfer to keratinocytes. J. Cell Sci., 115, 1441–1451. Singh, S.K., Nizard, C., Kurfurst, R., Bonte, F., Schnebert, S., and Tobin, D.J. (2008) The silver locus product (Silv/ gp100/Pmel17) as a new tool for the analysis of melanosome transfer in human melanocyte–keratinocyte co-culture. Exp. Dermatol., 17, 418–426. Bohil, A.B., Robertson, B.W., and Cheney, R.E. (2006) Myosin-X is a molecular motor that functions in ﬁlopodia formation. Proc. Natl. Acad. Sci. USA, 103, 12411–12416. Watanabe, T.M., Tokuo, H., Gonda, K., Higuchi, H., and Ikebe, M. (2010) Myosin-X induces ﬁlopodia by multiple

98

99

100

101

102

103

104

105

106

elongation mechanism. J. Biol. Chem., 285, 19605–19614. Cox, D., Berg, J.S., Cammer, M., Chinegwundoh, J.O., Dale, B.M., Cheney, R.E., and Greenberg, S. (2002) Myosin X is a downstream effector of PI3K during phagocytosis. Nat. Cell Biol., 4, 469–477. Seiberg, M. (2001) Keratinocyte– melanocyte interactions during melanosome transfer. Pigment Cell Res., 14, 236–242. Santulli, R.J., Derian, C.K., Darrow, A.L., Tomko, K.A., Eckardt, A.J., Seiberg, M., Scarborough, R.M., and AndradeGordon, P. (1995) Evidence for the presence of a protease-activated receptor distinct from the thrombin receptor in human keratinocytes. Proc. Natl. Acad. Sci. USA, 92, 9151–9155. Seiberg, M., Paine, C., Sharlow, E., Andrade-Gordon, P., Costanzo, M., Eisinger, M., and Shapiro, S.S. (2000) The protease-activated receptor 2 regulates pigmentation via keratinocyte– melanocyte interactions. Exp. Cell Res., 254, 25–32. Seiberg, M., Paine, C., Sharlow, E., Andrade-Gordon, P., Costanzo, M., Eisinger, M., and Shapiro, S.S. (2000) Inhibition of melanosome transfer results in skin lightening. J. Invest. Dermatol., 115, 162–167. Sharlow, E.R., Paine, C.S., Babiarz, L., Eisinger, M., Shapiro, S., and Seiberg, M. (2000) The protease-activated receptor-2 upregulates keratinocyte phagocytosis. J. Cell Sci., 113, 3093–3101. Scott, G., Deng, A., Rodriguez-Burford, C., Seiberg, M., Han, R., Babiarz, L., Grizzle, W., Bell, W., and Pentland, A. (2001) Protease-activated receptor 2, a receptor involved in melanosome transfer, is upregulated in human skin by ultraviolet irradiation. J. Invest. Dermatol., 117, 1412–1420. Babiarz-Magee, L., Chen, N., Seiberg, M., and Lin, C.B. (2004) The expression and activation of protease-activated receptor-2 correlate with skin color. Pigment Cell Res., 17, 241–251. Cardinali, G., Ceccarelli, S., Kovacs, D., Aspite, N., Lotti, L.V., Torrisi, M.R., and Picardo, M. (2005) Keratinocyte growth

321

322

10 Transport and Distribution of Melanosomes

107

108

109

110

111

112

113

114

factor promotes melanosome transfer to keratinocytes. J. Invest. Dermatol., 125, 1190–1199. Scott, G., Leopardi, S., Parker, L., Babiarz, L., Seiberg, M., and Han, R. (2003) The proteinase-activated receptor-2 mediates phagocytosis in a Rho-dependent manner in human keratinocytes. J. Invest. Dermatol., 121, 529–541. Tang, A., Eller, M.S., Hara, M., Yaar, M., Hirohashi, S., and Gilchrest, B.A. (1994) E-cadherin is the major mediator of human melanocyte adhesion to keratinocytes in vitro. J. Cell Sci., 107, 983–992. Goh, B., Kumarasinghe, P., and Lee, Y. (2005) Loss of melanosome transfer accounts for gluttate leucoderma in Darier’s disease: electron microscopic ﬁndings. Pigment Cell Res., 18, 48. Minwalla, L., Zhao, Y., Cornelius, J., Babcock, G.F., Wickett, R.R., Le Poole, I.C., and Boissy, R.E. (2001) Inhibition of melanosome transfer from melanocytes to keratinocytes by lectins and neoglycoproteins in an in vitro model system. Pigment Cell Res., 14, 185–194. Greatens, A., Hakozaki, T., Koshoffer, A., Epstein, H., Schwemberger, S., Babcock, G., Bissett, D., Takiwaki, H., Arase, S., Wickett, R.R., and Boissy, R.E. (2005) Effective inhibition of melanosome transfer to keratinocytes by lectins and niacinamide is reversible. Exp. Dermatol., 14, 498–508. Cerdan, D., Redziniak, G., Bourgeois, C.A., Monsigny, M., and Kieda, C. (1992) C32 human melanoma cell endogenous lectins: characterization and implication in vesicle-mediated melanin transfer to keratinocytes. Exp. Cell Res., 203, 164–173. Thong, H.Y., Jee, S.H., Sun, C.C., and Boissy, R.E. (2003) The patterns of melanosome distribution in keratinocytes of human skin as one determining factor of skin colour. Br. J. Dermatol., 149, 498–505. Minwalla, L., Zhao, Y., Le Poole, I.C., Wickett, R.R., and Boissy, R.E. (2001) Keratinocytes play a role in regulating distribution patterns of recipient

115

116

117

118

119

120

121

122

melanosomes in vitro. J. Invest. Dermatol., 117, 341–347. Yoshida, Y., Hachiya, A., Sriwiriyanont, P., Ohuchi, A., Kitahara, T., Takema, Y., Visscher, M.O., and Boissy, R.E. (2007) Functional analysis of keratinocytes in skin color using a human skin substitute model composed of cells derived from different skin pigmentation types. FASEB J., 21, 2829–2839. Weiner, L., Han, R., Scicchitano, B.M., Li, J., Hasegawa, K., Grossi, M., Lee, D., and Brissette, J. (2007) Dedicated epithelial recipient cells determine pigmentation patterns. Cell, 130, 932–942. Duval, C., Regnier, M., and Schmidt, R. (2001) Distinct melanogenic response of human melanocytes in mono-culture, in co-culture with keratinocytes and in reconstructed epidermis, to UV exposure. Pigment Cell Res., 14, 348–355. Kobayashi, N., Nakagawa, A., Muramatsu, T., Yamashina, Y., Shirai, T., Hashimoto, M.W., Ishigaki, Y., Ohnishi, T., and Mori, T. (1998) Supranuclear melanin caps reduce ultraviolet induced DNA photoproducts in human epidermis. J. Invest. Dermatol., 110, 806–810. Gibbs, S., Murli, S., De Boer, G., Mulder, A., Mommaas, A.M., and Ponec, M. (2000) Melanosome capping of keratinocytes in pigmented reconstructed epidermis – effect of ultraviolet radiation and 3-isobutyl-1methyl-xanthine on melanogenesis. Pigment Cell Res., 13, 458–466. Byers, H.R., Maheshwary, S., Amodeo, D.M., and Dykstra, S.G. (2003) Role of cytoplasmic dynein in perinuclear aggregation of phagocytosed melanosomes and supranuclear melanin cap formation in human keratinocytes. J. Invest. Dermatol., 121, 813–820. Byers, H.R., Dykstra, S.G., and Boissel, S.J. (2007) Requirement of dynactin p150Glued subunit for the functional integrity of the keratinocyte microparasol. J. Invest. Dermatol., 127, 1736–1744. Borovansky, J. and Elleder, M. (2003) Melanosome degradation: fact or ﬁction. Pigment Cell Res., 16, 280–286.

323

11 Genetics of Melanosome Structure and Function Vincent J. Hearing

11.1 Introduction

In mammals, melanins are synthesized within specialized membrane-bound organelles termed melanosomes that are exclusively produced in melanocytes, as detailed in other chapters throughout this book. The eventual colors and patterns generated, be they in the hair, skin, and/or eye, depend on a complex series of physiologically regulated events. Those begin during the development of melanoblasts, continue throughout the differentiation and function of melanocytes in their destination tissues, and culminate with the synthesis and distribution of melanin pigments in those tissues to produce the wide variety of colors in animals that we are familiar with. Each of those steps is regulated by a number of distinct genes that control those processes in very sensitive and speciﬁc manners. Currently, more than 250 genes are known to affect mammalian pigmentation either directly or indirectly (see http://www.espcr.org/micemut/ for the current list) and even minor perturbations in the functions of their encoded proteins can have dramatic consequences on eventual color, and often on other cellular and tissue processes that depend in some context on melanocyte function. There has been a recent surge in interest in studying the genetics of pigmentation, not only because of the important functions of melanins (e.g., camouﬂage and photoprotection), but also because the highly visible colors that are produced provide an easy visible output assay to characterize the full complement of processes (e.g., intracellular signaling and transcription factors) that regulate various cellular functions. This chapter focuses on genes that regulate melanosome structure and function; however, to put those in context, will begin with a general review of genes important to the development, growth, survival, and differentiation of melanocytes – the cells that produce the melanins and melanosomes. It is important to keep in mind that colors (and their patterns) ultimately depend upon the presence of functional melanocytes in any given area (e.g., consider the black and white stripes on a zebra), and also how much and what types of melanin they are producing (e.g., consider the black versus orange stripes of a tiger and even the decreased pigmentation in ventral areas). It is the sum of the development, migration, proliferation, Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

324

11 Genetics of Melanosome Structure and Function

Figure 11.1

Processes critical to getting functional melanocytes in place.

survival, and even regulated function of melanocytes in various areas of tissues that gives rise to the wide diversity of pigmentation patterns seen in nature. This is not merely a cosmetic issue, since melanins in lower species play critical roles in survival, such as camouﬂage for predator and prey, and in regulating body temperature in amphibians and snakes; in man, they play important roles in the absorption of toxic materials, in the scavenging of free radicals, and in photoprotection against UV damage. More complete reference lists and further information can be found in [1–3], and a schematic summarizing these processes is shown in Figure 11.1.

11.2 Genes Involved in Melanoblast Development, Migration, and Speciﬁcation

Melanocytes that eventually populate the skin and other locations in the body begin as precursor cells termed melanoblasts. In vertebrates, all pigment cells (except for those destined for the retinal pigment epithelium of the eye) are initially derived from the dorsal neural tube during embryonic development. To get to their eventual destinations in the body, cues must be provided to these precursor cells to begin migrating from the neural tube along deﬁned pathways that take them to their distant destinations, they must continue to migrate along those pathways

11.3 Genes Involved in Melanocyte Differentiation, Survival, and Proliferation

until they reach their targeted destination, and then obviously they must stop migrating and begin to transition into functional melanocytes (the latter process being termed “speciﬁcation”). Each of those steps is under the continual control of many independent factors (including intrinsic and environmental factors) and the overall timing of these processes is critical to achieve the adult pigmentation patterns in a given normal individual of any species. The genes that function at this level encode, among other things, transcription factors (e.g., PAX3, SOX10, and MITF), receptors and their ligands (e.g., EDNRB/EDN3, KIT/KITL, and FZD4/WNT3a), and other factors important to the start/continue/stop signals for migration, speciﬁcation, etc (e.g., ADAMTS20, MCOLN3, and ITGB1). Mutations in those genes and/or disruptions in their functions frequently perturb normal development and lead to developmental diseases that not only affect pigmentation, but in many cases also affect tissues that depend in some fashion on the functions of cells derived from the neural tube. Regarding effects on pigmentation, those diseases typically lead to white-spotted areas where melanoblasts failed to develop and migrate to (typically on the forehead and abdomen) that are stable (i.e., present at birth and remain through adulthood). Examples of such developmental diseases include Waardenburg syndrome, Hirschsprung’s disease, and piebaldism. It is beyond the scope of this chapter to provide details of the genes and processes involved or the pigmentary diseases that result, and interested readers are referred to recent reviews for pertinent details [4–11] and also to relevant chapters in this book. A summary of genes involved in melanoblast development, migration, and speciﬁcation, along with the functions of their encoded proteins and associated diseases where known is provided in Table 11.1.

11.3 Genes Involved in Melanocyte Differentiation, Survival, and Proliferation

Once melanoblasts have arrived at their ﬁnal destinations in the tissues, they must then differentiate to functional melanocytes, they must survive, and then must proliferate to populate the tissue to achieve the correct eventual density of melanocytes required for normal function. Again, a relatively large number of genes are involved in those processes and disruption in any of them leads to decreased melanocyte function in the adult. Again, the genes involved in these processes encode, among other things, transcription factors (e.g., MITF and FOXD3), receptors and their ligands (e.g., EDNRB/EDN3 and KIT/KITL), and other factors important to the signals that regulate proliferation, survival, and so on (e.g., BCL2, SEMA3C, and NLRP1). Note that many of these critical genes function at multiple stages, ranging from developmental processes (as noted in the previous section) through differentiation processes (as discussed below). Genes that are critical to many different levels that affect pigmentation include transcription factors (such as MITF and SOX10) and receptor/ligand pairs (such as MC1R/POMC and EDNRB/EDN3). Regarding effects on pigmentation, mutations in those genes typically lead to hypopigmented areas where melanoblasts failed to differentiate or

325

326

11 Genetics of Melanosome Structure and Function Genes and diseases involved with melanoblast development, migration, and speciﬁcation.

Table 11.1

Gene

Encoded protein

Function

Associated disease/ condition

OMIM #

PAX3

splotch

transcription factor

Waardenburg syndrome types 1, 3

193500

SOX9

Sry box gene 9

transcription factor

unknown

SOX10

Sry box gene 10

transcription factor

Waardenburg–Shah syndrome

277580

SNAI2

snail homolog 2

transcription factor

Waardenburg syndrome type 2

193510

MITF

microphthalmia

transcription factor

Waardenburg syndrome type 2

193510

PAX6

paired box gene 6

transcription factor

aniridia

106210

LEF1

lymphoid binding factor 1

transcription factor

sebaceous adenoma

153245

TCFAP2A

AP2α

transcription factor

unknown

FOXD3

forkhead box D3

transcription factor

vitiligo

607836

EDNRB

endothelin B receptor

receptor

Hirschsprung’s disease type 2

600155

EDN3

endothelin 3

EDNRB ligand

Waardenburg–Shah syndrome

277580

EDN1

endothelin 1

EDNRB ligand

unknown

FGFR2

ﬁbroblast growth factor receptor 2

receptor

Crouzon syndrome

FGF

ﬁbroblast growth factor

FGFR2 ligand

unknown

KIT

Kit oncogene

receptor

piebaldism

KITL/SCF

stem cell factor

KIT ligand

unknown

FZD4

frizzled homolog 4

receptor

exudative vitreoretinopathy

WNT1

wingless related 1

FZD4 ligand

unknown

WNT3A

wingless related 3a

FZD4 ligand

unknown

123500

172800

133780

11.4 Genes Involved in Regulating Melanocyte Function Table 11.1 (Continued)

Gene

Encoded protein

Function

Associated disease/ condition

MET

Met oncogene

receptor

unknown

HGF

hepatocyte growth factor

MET ligand

unknown

EGFR

epidermal growth factor receptor

receptor

unknown

ITGB1

integrin β1

receptor

unknown

MCOLN3

mucolipin 3

cation channel

unknown

ADAMTS20

disintegrin protease 20

metalloprotease

unknown

OMIM #

survive that are progressive (i.e., pigmentation may be normal at birth, but become more severe with age). Examples of such diseases include vitiligo and retinitis pigmentosum. Again, it is beyond the scope of this chapter to go into further detail about these processes and how the relevant genes are involved, and interested readers are encouraged to look at recent review articles [12–18] and also the relevant chapters in this book. A partial list of genes involved with these processes at this level and diseases associated with their dysfunction is shown in Table 11.2.

11.4 Genes Involved in Regulating Melanocyte Function

Once functional melanocytes have been established and become active in the appropriate locations and densities in various tissues, the next level of control involves the regulation of their functions (i.e., their phenotypic characteristics such as their ability to produce melanin and melanosomes, and what types of melanins are produced and how much). Interestingly, many of the same genes discussed above (and their encoded products) are involved at this level of regulation. Obviously, mutations in those multilevel genes affect pigmentation in developmental stages initially and are associated with developmental diseases as noted above, but the functions of their encoded proteins also regulate melanocyte function, and thus modulation of their expression and function in adult tissues leads to often dramatic changes in pigmentation. This list of genes is far from complete at this point and the bulk of as yet uncloned pigment genes (see http://www.espcr.org/ micemut/ for an updated list) no doubt includes a large number of genes that operate at this level, such as genes that regulated constitutive colors in the skin, hair, and eyes or that regulate environmental responses such as those modulated by UV exposure, as summarized in Table 11.3.

327

328

11 Genetics of Melanosome Structure and Function Genes and diseases involved with melanocyte differentiation, survival and proliferation.

Table 11.2

Gene

Encoded protein

Function

Associated disease/condition

OMIM #

MITF

microphthalmia

transcription factor

Waardenburg syndrome type 2

193510

FOXD3

forkhead box D3

transcription factor

vitiligo

607836

EDNRB

endothelin B receptor

receptor

Hirschsprung’s disease type 2

600155

EDN3

endothelin 3

EDNRB ligand

Waardenburg– Shah syndrome

277580

EDN1

endothelin 1

EDNRB ligand

unknown

KIT

Kit oncogene

receptor

piebaldism

KITL/SCF

stem cell factor

KIT ligand

unknown

BCL2

B cell leukemia 2

inhibit apoptosis

follicular lymphoma

GNAQ

GNA subunit Gaq

G-protein-coupled receptor signaling

unknown

GNA11

GNA subunit Ga11

G-protein-coupled receptor signaling

unknown

RB1

retinoblastoma 1

growth inhibitor

retinoblastoma

SEMA3C

semaphorin 3c

signaling factor

unknown

SEMA4A

semaphorin 4a

signaling factor

retinitis pigmentosum

610282

NLRP1

NLR protein 1

immune regulator

vitiligo

606579

172800

151430

180200

Visible pigmentation is regulated physiologically under normal conditions, such as to provide the wide range of constitutive skin, hair, and eye colors in various races and individuals, and also to respond to various stresses, such as exposure to UV radiation and/or injury. Effects on visible pigmentation can be short-term (e.g., temporary increases (tans) due to UV exposure) and/or more permanent (e.g., age spots and postinﬂammatory hyperpigmentation). Again, transcription factors and receptor/ligand interactions play major roles in regulating these responses, but a number of other factors, such as membrane-bound transporters and proteolytic components, also have important functions in this respect, as noted in Table 11.3. The following is a brief consideration of genes and processes involved with the regulation of hair, skin, and eye color under “normal” physiological conditions,

11.4 Genes Involved in Regulating Melanocyte Function Table 11.3 Genes and diseases involved with regulating melanocyte function.

Gene

Encoded protein

Function

Associated disease/condition

MITF

microphthalmia

SOX9

Sry box gene 9

SOX2

Sry box gene 2

hair, skin, and eye color skin phenotype, response to UV unknown

SOX10

Sry box gene 10

SOX18

Sry box gene 18

PAX6

paired box gene 6

FOXD3

forkhead box D3

MC1R

KIT

melanocortin 1 receptor proopiomelanocortin agouti signaling protein Kit oncogene

transcription factor transcription factor transcription factor transcription factor transcription factor transcription factor transcription factor receptor MC1R ligand MC1R antagonist receptor

KITL

stem cell factor

KIT ligand

EDNRB

endothelin B receptor

receptor

EDN3 EDN1

endothelin 3 endothelin 1

EDNRB ligand EDNRB ligand

MET

receptor growth factor

UV-A melanosis

DKK1

hepatocyte growth factor receptor granulocyte macrophage colony-stimulating factor dickkopf 1

hair and skin color hair, skin, and eye color UV-B melanosis, lentigo senilis UV-B melanosis, lentigo senilis, hair color UV-B melanosis, lentigo senilis unknown UV-B melanosis, lentigo senilis UV-A melanosis

Wnt inhibitor

NRG1

neuregulin 1

P

P protein

TYR

tyrosinase

TYRP1

tyrosinase-related protein 1

Erb receptor ligand transport of ??? tyrosinase enzyme tyrosinase stability

skin color and thickness skin color

POMC ASP

GM-CSF

OMIM #

unknown unknown aniridia, eye color

607108

vitiligo

607836

hair and skin color

eye and skin color skin color eye and hair color

(Continued)

329

330

11 Genetics of Melanosome Structure and Function Table 11.3

(Continued)

Gene

Encoded protein

Function

Associated disease/condition

SLC45A2

solute carrier 45 A2

eye and skin color

SLC7A11

solute carrier 7 A11

SLC24A5

solute carrier 24 A5

ATP7A

ATPase 7α

ATP7B

ATPase 7β

CTNS

cystinosin

ATOX1

antioxidant protein 1

ATRN MGRN1 DEFB103A

mahagony mahogunin ring ﬁnger 1 β-defensin 3

PMEL17

gp100/silver

RAB7 IL1

Ras-associated protein 7 interleukin-1

TNFA

tumor necrosis factor

IFNG

interferon-γ

transport of ??? cysteine transport calcium transport copper transport copper transport cysteine transport copper transport eu- versus pheoE3 ubiquitin ligase MC1R activation melanosome structure protein trafﬁcking inﬂammatory cytokine inﬂammatory cytokine inﬂammatory cytokine

OMIM #

unknown skin color Menkes disease

309400

Wilson disease

277900

cystinosis

219800

unknown unknown unknown unknown hair color unknown postinﬂammatory hyperpigmentation lentigo senilis postinﬂammatory hyperpigmentation

followed by those involved with hypopigmentation and, ﬁnally, by those involved with hyperpigmentation. For more details of these regulatory processes, and listings of the original references, see several recent reviews [11, 19–29]. See Figure 11.2. 11.4.1 Regulation of Constitutive Skin, Hair, and Eye Color

In this section, the wide variations in the colors of “normal” individuals (i.e., those without pigmentary diseases as covered in Sections 11.5–11.7) is discussed. Genes that elicit various pigmentary diseases obviously play critical roles in regulating skin, hair, and eye color that supersede the actions of genes controlling normal variation and responses to the environment. Interestingly, although the biochemi-

11.4 Genes Involved in Regulating Melanocyte Function

Figure 11.2 Factors and signaling pathways regulating melanocyte function. (Adapted from

[26].)

cal processes involved in producing melanins are essentially identical in various tissues and similar regulatory steps are involved, independent and distinct sets of genes seem to play roles in regulating the constitutive colors of the hair, skin, and eyes. This probably results from the fact that the fate of melanins and melanosomes produced varies among those tissues: in the skin, melanosomes are transferred from melanocytes to keratinocytes and are gradually moved up towards the stratum corneum (over 3–4 weeks) and are lost by desquamation. In contrast, in the hair bulbs, melanosomes are transferred to keratinocytes in the growing hair shaft and are very slowly (over the course of months) taken upwards to emerge from the surface of the skin, whereas in the eye, melanosomes produced by the choroid and retina remain within those cells and are not transferred, thus staying dormant for years. The regulation of hair color in normal individuals is probably the best characterized and depends almost solely on MC1R function (wild-type MC1R is regulated by the agonist MSH/POMC and the antagonist ASIP, and by various modiﬁers that modulate its ligand binding ability). Disruptions in MC1R function are associated with the red hair/fair skin phenotype; MC1R has 30 known variant alleles, about three of which cause virtually complete losses in receptor activity and cause red hair [30–33]. The genes encoding POMC and ASIP also play important roles in regulating hair color (see recent reviews by [32, 34–38]). Several other genes also regulate hair color, at least based on the phenotype of mutant

331

332

11 Genetics of Melanosome Structure and Function

mice, including the KITL locus (called steel in mice) and PMEL17 (called silver in mice) as reviewed in [33]. The regulation of eye color is also relatively well characterized, and is determined principally by three loci – P, MITF, and PAX6 (reviewed in [39]). It has been estimated that around 75% of human eye color is controlled by the P locus, which encodes a transmembrane protein that has an as yet unknown function, but is somehow involved in regulating tyrosinase function in melanosomes [40]. In contrast, skin color is by far the most complex phenotype and is regulated by a wide variety of genes. Skin color is regulated by loci that regulate eye and hair color, but is also dependent on other genes expressed by melanocytes, including SLC24A5 and SLC45A2. In addition to those genes, factors secreted by other cells in the skin dramatically affect skin pigmentation, most notably two factors secreted by ﬁbroblasts in the underlying dermis: DKK1, which inhibits pigmentation in palmoplantar skin compared to trunk skin [26, 41], and NRG1, which modulates the darker skin phenotypes [42]. Several recent reviews have been written about factors that regulate skin pigmentation and the mechanisms by which that occurs [25, 26, 43]. 11.4.2 Hypopigmentation

Decreased pigmentation in adult tissues can result from the death of melanocytes in localized areas, such as occurs in white macules (termed guttate leukoderma) in the skin [44, 45] and in occupational vitiligo [46]. It can also result from the reduced function of components of the melanogenic scheme that decrease melanin production by tyrosinase, such as decreased copper transport by ATP7A or ATP7B (copper being an essential cofactor required for tyrosinase catalytic function), decreased uptake of tyrosine or phenylalanine (precursors of melanins), and increased uptake of cysteine (which leads to a less pigmented type of melanin known as pheomelanin). As mentioned above, vitiligo is due to the destruction of melanocytes in the skin, typically a progressive condition, and an ever-increasing number of genes have been associated with that condition, the majority of them involved in immune function, suggesting that melanocytes in vitiliginous skin may be targeted and destroyed by the immune system [47–49]. 11.4.3 Hyperpigmentation

Increased pigmentation is a more common phenomenon (perhaps because it is much more visible) and is a typical response to a wide variety of environmental stresses [25]. The most well-known pigmentary response is the tanning response to UV radiation, and in fact there are four distinct types of tanning responses, each due to different mechanisms, as recently reviewed [24, 50]. Large numbers of genes respond to different types of UV radiation, including those expressed by ﬁbroblasts, keratinocytes, and melanocytes [51], which as an end result elicit increased visible pigmentation of the skin, which can last for hours, days, weeks,

11.5 Genes Involved in the Biogenesis of Melanosomes and Other Lysosome-Related Organelles

and even years. The genes involved in UV responses are numerous, but major ones include familiar receptor/ligand pairs (e.g., EDNRB/EDN3 and KIT/ KITL) and transcription factors (e.g., MITF and SOX9). Other types of hyperpigmentation include postinﬂammatory hyperpigmentation, age spots (lentigo senilis), and melasma. Postinﬂammatory hyperpigmentation typically occurs in areas that have been stressed by injury, and the immune mediators released in those areas stimulate melanocyte differentiation and function (e.g., interleukin-1 and interferon-γ). Age spots are thought to result from long-term damage due to UV exposure and many of the same receptor/ligand pairs have been associated with the increased hyperpigmentation of these lesions (e.g., EDNRB/EDN1 and KIT/KITL).

11.5 Genes Involved in the Biogenesis of Melanosomes and Other Lysosome-Related Organelles

As discussed in detail in Chapter 9, melanosomes belong to the family of lysosome-related organelles (LROs), which are essentially produced by the same cellular mechanism that leads to the formation of lysosomes in all cell types, but with speciﬁc constituents added in speciﬁc types of cells to make the functions of those organelles unique to that cell type. The family of LROs includes melanosomes, lysosomes, platelet-dense bodies, synaptosomes, and so on. Mutations in genes that affect the general process of LRO biogenesis cause disruptions of organelle function in many types of cells in various tissues that depend on the functions of those LROs. Known factors (and their encoding genes) that function at this level are listed in Table 11.4 and further details can also be found in several recent review articles [52–57]. The effects in patients with Hermansky–Pudlak syndrome are pleiotropic, affecting bleeding/clotting (due to effects on plateletdense bodies), immune functions (due to effects on lysosomes), and pigmentation (due to effects on melanosomes). Studies of pigment genes have led to remarkable advances in our understanding of the mechanisms of organelle biogenesis simply because the resulting effects on pigmentation are highly visible, as are molecular approaches to rescue gene function (and pigmentation). Most genes involved at this level have functions in the processing and sorting (trafﬁcking) of organelle components; for example, on proteins that have protein recognition and docking duties as well as subcellular complexes (such as LBOC1, BLOC2, etc.) that are involved in the trafﬁcking of organelle components. Characterization of the functions of proteins encoded by pigment genes that regulate pigmentation at this level have led so far to the identiﬁcation of eight types of Hermansky– Pudlak syndrome and one type of Chediak–Higashi syndrome, but there are many other cloned genes in this category with as yet unknown functions and no doubt quite a few that have not yet been cloned but will also belong in this category. Thus, many more subtypes of these diseases are expected to be deﬁned in the future.

333

334

11 Genetics of Melanosome Structure and Function Table 11.4

Genes and diseases involved in the biogenesis of melanosomes and other LROs.

Gene

Encoded protein

Function

Associated disease/ condition

OMIM #

HPS1

HPS1

LRO biogenesis

Hermansky–Pudlak syndrome type 1

604982

HPS2

adaptor protein β3

LRO biogenesis

Hermansky–Pudlak syndrome type 2

603401

HPS3

HPS3

LRO biogenesis

Hermansky–Pudlak syndrome type 3

606118

HPS4

HPS4

LRO biogenesis

Hermansky–Pudlak syndrome type 4

606682

HPS5

HPS5/BLOC2

LRO biogenesis

Hermansky–Pudlak syndrome type 5

607521

HPS6

HPS6/BLOC2

LRO biogenesis

Hermansky–Pudlak syndrome type 6

607522

HPS7

DTNBP1/dysbindin

LRO biogenesis

Hermansky–Pudlak syndrome type 7

203300

HPS8

HPS8/BLOC1S3

LRO biogenesis

Hermansky–Pudlak syndrome type 8

609762

LYST

LYST

LRO biogenesis

Chediak–Higashi syndrome

214500

PLDN

pallidin

Vesicle docking

unknown

VPS33A

vacuolar protein sorting 33A

LRO trafﬁcking

unknown

RAB38

Ras-associated protein 38

TYRP1 trafﬁcking

unknown

RABGGTA

gunmetal (Rab geranylgeranyl transferase α subunit)

LRO biogenesis

unknown

AP3D1

adapter protein δ1

LRO trafﬁcking

unknown

MUTED

Txndc5/BLOC1

LRO biogenesis

unknown

11.6 Genes Involved in Melanin Production

To date, about 12 genes have been cloned and characterized whose encoded proteins function in melanosomes, and thus are speciﬁc to melanocytes and pigmentation. Those proteins regulate the type and amount of melanins produced, and thus play important roles in regulating the constitutive pigmentation of tissues

11.6 Genes Involved in Melanin Production Table 11.5 Genes and diseases involved with melanin production.

Gene

Encoded protein

Function

Associated disease/ condition

OMIM #

TYR

tyrosinase

melanogenic enzyme

oculocutaneous albinism type 1

203100

P

P protein

unknown

oculocutaneous albinism type 2

203200

TYRP1

tyrosinaserelated protein 1

tyrosinase stability

oculocutaneous albinism type 3

203290

SLC45A2

solute carrier 45 A2/MATP

unknown

oculocutaneous albinism type 4

606574

GPR143

OA1

melanosome structure

ocular albinism type 1

300500

DCT

dopachrome tautomerase

melanogenic enzyme

unknown

PMEL17

gp100/silver

melanosome structure

unknown

GPNMB

GPNMB

melanosome component

pigment dispersion syndrome

SLC24A5

solute carrier 24 A5

calcium transport

unknown

OSTM1

gray-lethal

pheomelanin synthesis

osteopetrosis

259700

PAH

phenylalanine hydroxylase

tyrosine synthesis

phenylketonuria

261600

MART1

melanoma antigen 1

melanosome structure

unknown

600510

and responses to the environment and other regulatory factors, as summarized in Table 11.5. Disruptions in the functions of those proteins (e.g., by mutations in their encoding genes) lead to pigment-speciﬁc diseases that typically are grouped in the category of oculocutaneous albinism (OCA). Thus far, four types of OCA have been identiﬁed, all of which affect in one way or another, the function of tyrosinase, the critical gene in melanin production (see Chapters 4 and 5 for more details). OCA1, the most severe type, results from mutations in the gene encoding tyrosinase, the critical gene required for melanin biosynthesis, as might be expected. However, interestingly, OCA2 and OCA4 are almost as severe phenotypes and result from disruptions of the P and MATP genes, respectively, which are critical for tyrosinase trafﬁcking to melanosomes, which is of course essential to its function. OCA3 is a less severe form of OCA that results from disruptions

335

336

11 Genetics of Melanosome Structure and Function

in TYRP1, a protein that acts as a chaperone to stabilize tyrosinase and ensure its delivery to melanosomes. However, TYRP1 is not absolutely required for tyrosinase function and some tyrosinase is delivered to melanosomes, hence the phenotype of OCA3 is relatively mild. A number of other genes encoding melanosome-speciﬁc proteins have been cloned, but have not yet associated with a pigmentary disease (e.g., DCT and PMEL17); it is probable that the phenotypes of those diseases are even milder than OCA3, but eventually they may be associated with novel types of OCA. The functions of proteins in this group include those with enzymatic functions involved in synthesizing various types of melanins (TYR, TYRP1, and DCT), those with structural functions that are essential to the unique structure of melanosomes (PMEL17 and MART1), proteins involved with the trafﬁcking of those proteins to melanosomes (P and SLC45A2), and proteins that have other functions, such as substrate pumps to deliver melanogenic substrates into the membrane-bound melanosomes (e.g., SLC24A5 and PAH), or that regulate other parameters important to organelle function or melanin synthesis, such as pH. Interested readers are referred to the following reviews for further details and original references [29, 33, 58–64].

11.7 Genes Involved in Melanosome Movement, Transfer, and Distribution

As a ﬁnal consideration of genes that affect pigmentation, the eventual distribution of melanin in the tissues is equally critically important to visible color, and that is, determined not only by the production and distribution of melanosomes within melanocytes, but also (in the case of the skin and hair) their transfer to neighboring keratinocytes and eventual movement towards the surface of the skin or the emerging hair shaft. Genes and factors regulating those later events have not yet been characterized well, at least from the molecular and genetic point of view. Since the three processes that function at this level (movement, transfer, and distribution) involve different subsets of cells (melanocytes only, melanocytes and keratinocytes, and keratinocytes only, respectively) distinct types of genes are involved, and those cloned and characterized to date are listed in Table 11.6. 11.7.1 Movement

As melanin is synthesized and deposited within melanosomes, they are usually moving from the perinuclear areas of the melanocytes to the peripheral dendritic areas. This intracellular movement of the granules and accumulation in the tips of dendrites is essential for their eventual transfer to neighboring cells, and several motor proteins have been shown to be involved in those dynamics, as reviewed recently in [65–73]. When the functions of any of those motor proteins in the melanosome transport complex (RAB27A, MLPH, and/or MYO5A) are disrupted, the melanosomes accumulate in the perinuclear area and are thus unable to be

11.7 Genes Involved in Melanosome Movement, Transfer, and Distribution Table 11.6 Genes and diseases involved with melanosome movement, transfer and distribution.

Gene

Encoded protein

Function

Associated disease/ condition

OMIM #

RAB27A

Ras-associated protein 27a

melanosome transport

Griscelli syndrome

214450

MLPH

melanophilin

melanosome transport

Griscelli syndrome

214450

MYO5A

myosin type Va

melanosome transport

Griscelli syndrome

214450

MYO7A

myosin type VIIa

melanosome transport

Usher syndrome type 1b

276903

MREG

melanoregulin

melanosome transport

unknown

PAR2

protease-activated receptor type 2

melanosome transfer

unknown

KGF

keratinocyte growth factor

melanosome transfer

unknown

transferred to other neighboring cells, which results in a dramatically hypopigmented phenotype, called Griscelli syndrome in humans. 11.7.2 Transfer

Once melanin granules are positioned in the tips of the dendrites of melanocytes and are ready to be transferred, they are gradually transferred and accumulate in neighboring keratinocytes, although the exact mechanism of the process remains controversial. The parameters involved have been recently reviewed [74–77] and are discussed in Chapter 10. The sum of the evidence suggests that multiple intercellular processes are involved in the transfer, and that factors in melanocytes and also in keratinocytes are involved in coregulating the process. 11.7.3 Distribution

The eventual distribution of melanins in upper layers of the epidermis and in the emerging hair shaft is responsible for most of the visible pigmentation, yet little is known about how that process is regulated, and at this point, no pigment genes have been cloned and characterized that regulate that process. Certainly that

337

338

11 Genetics of Melanosome Structure and Function

distribution can be dramatically affected (e.g., in light versus dark skin and by exposure to UV as part of the tanning process). Interested readers are referred to several recent reviews on these topics [24–26, 78–82].

11.8 Conclusions

The synthesis, deposition, and distribution of melanin pigments play critical roles not only in the pigmentation of various tissues, but also in their functions. These processes are under the control of a wide range of genes that act at various levels during development, differentiation, and regulation of melanocyte function. The impact of these processes is seen easily in human skin, where melanin content plays important roles in social interactions, sexual attraction, and protection against skin cancers, among other parameters. The function(s) of melanins is less evident in other tissues and organs where melanocytes are present in less abundance, yet they are important during development and/or differentiation, in tissues such as the brain, inner ear, heart, and so on, as addressed in Chapters 2, 7, and 8, and also in several recent reviews [83, 84].

References 1 Nordlund, J.J., Boissy, R.E., Hearing, V.J.,

2

3

4

5

Oetting, W.S., King, R.A., and Ortonne, J.P. (2006) The Pigmentary System: Physiology and Pathophysiology, 2nd edn, Blackwell, Malden, MA. King, R.A., Oetting, W.S., Summers, C.G., Creel, D.J., and Hearing, V.J. (2006) Abnormalities of pigmentation, in Emery and Rimoin’s Principles and Practice of Medical Genetics (eds D.L. Rimoin, J.M. Connor, R.E. Pyeritz, and B.R. Korf), Churchill Livingstone, New York, pp. 3380–3428. Hearing, V.J. and Leong, S.P.L. (2005) From Melanocyte to Melanoma: The Progression to Malignancy, Humana Press, Totowa, NJ. Saito, H., Yasumoto, K., Takeda, K., Takahashi, K., Yamamoto, H., and Shibahara, S. (2003) Microphthalmiaassociated transcription factor in the Wnt signaling pathway. Pigment Cell Res., 16, 261–265. Baxter, L.L., Hou, L., Loftus, S.K., and Pavan, W.J. (2004) Spotlight on spotted mice: a review of white spotting mouse

6

7

8

9

10

11

mutants and associated human pigmentation disorders. Pigment Cell Res., 17, 215–224. Tomita, Y. and Suzuki, T. (2004) Genetics of pigmentary disorders. Am. J. Med. Genet. C, 131C, 75–81. Thomas, A.J. and Erickson, C.A. (2008) The making of a melanocyte: the speciﬁcation of melanoblasts from the neural crest. Pigment Cell Melanoma Res., 21, 598–610. Kelsh, R.N., Harris, M.L., Colanesi, S., and Erickson, C.A. (2009) Stripes and belly-spots – a review of pigment cell morphogenesis in vertebrates. Semin. Cell Dev. Biol., 20, 90–104. Medic, S. and Ziman, M. (2009) PAX3 across the spectrum: from melanoblast to melanoma. Crit. Rev. Biochem. Mol. Biol., 2009, 1–13. Ernfors, P. (2010) Cellular origin and developmental mechanisms during the formation of skin melanocytes. Exp. Cell Res., 316, 1397–1407. Vachtenheim, J. and Borovansky, J. (2010) “Transcription physiology” of

References

12

13

14

15

16

17

18

19

20

21

22

23

pigment formation in melanocytes: central role of MITF. Exp. Dermatol., 19, 617–627. Robinson, K.C. and Fisher, D.E. (2009) Speciﬁcation and loss of melanocyte stem cells. Semin. Cell Dev. Biol., 20, 111–116. Bedogni, B. and Powell, M.B. (2009) Hypoxia, melanocytes and melanoma – survival and tumor development in the permissive microenvironment of the skin. Pigment Cell Melanoma Res., 22, 166–174. Hornyak, T.J. (2006) The developmental biology of melanocytes and its application to understanding human congenital disorders of pigmentation. Adv. Dermatol., 22, 201–218. Levy, C., Khaled, M., and Fisher, D.E. (2006) MITF: master regulator of melanocyte development and melanoma oncogene. Trends Mol. Med., 12, 406–414. Passeron, T., Mantoux, F., and Ortonne, J.P. (2005) Genetic disorders of pigmentation. Clin. Dermatol., 23, 56–67. Vance, K.W. and Goding, C.R. (2004) The transcription network regulating melanocyte development and melanoma. Pigment Cell Res., 17, 318–325. Dupin, E. and Le Douarin, N.M. (2003) Development of melanocyte precursors from the vertebrate neural crest. Oncogene, 22, 3016–3023. Busca, R. and Ballotti, R. (2000) Cyclic AMP: a key messenger in the regulation of skin pigmentation. Pigment Cell Res., 13, 60–69. Imokawa, G. (2004) Autocrine and paracrine regulation of melanocytes in human skin and in pigmentary disorders. Pigment Cell Res., 17, 96–110. Slominski, A., Tobin, D.J., Shibahara, S., and Wortsman, J. (2004) Melanin pigmentation in mammalian skin and its hormonal regulation. Physiol. Rev., 84, 1155–1228. Slominski, A., Wortsman, J., Plonka, P.M., Schallreuter, K.U., Paus, R., and Tobin, D.J. (2005) Hair follicle pigmentation. J. Invest. Dermatol., 124, 13–21. García-Borrón, J.C., Sánchez-Laorden, B.L., and Jiménez-Cervantes, C. (2005) Melanocortin-1 receptor structure and functional regulation. Pigment Cell Res., 18, 393–410.

24 Miyamura, Y., Coelho, S.G., Wolber, R.,

25

26

27

28

29

30

31

32

33

34

35

Miller, S.A., Wakamatsu, K., Zmudzka, B.Z., Ito, S., Smuda, C., Passeron, T., Choi, W., Batzer, J., Yamaguchi, Y., Beer, J.Z., and Hearing, V.J. (2007) Regulation of human skin pigmentation and responses to ultraviolet radiation. Pigment Cell Res., 20, 2–13. Costin, G.E. and Hearing, V.J. (2007) Human skin pigmentation: melanocytes modulate skin color in response to stress. FASEB J., 21, 976–994. Yamaguchi, Y., Brenner, M., and Hearing, V.J. (2007) The regulation of skin pigmentation. J. Biol. Chem., 282, 27557–27561. Brenner, M. and Hearing, V.J. (2008) The protective role of melanin against UV damage in human skin. Photochem. Photobiol., 84, 539–549. Schallreuter, K.U., Kothari, S., Chavan, B., and Spencer, J.D. (2008) Regulation of melanogenesis – controversies and new concepts. Exp. Dermatol., 17, 395–404. Schiafﬁno, M.V. (2010) Signaling pathways in melanosome biogenesis and pathology. Int. J. Biochem. Cell Biol., 42, 1094–1104. Walker, W.P. and Gunn, T.M. (2010) Shades of meaning: the pigment-type switching system as a tool for discovery. Pigment Cell Melanoma Res., 23, 485–495. Walker, W.P. and Gunn, T.M. (2010) Piecing together the pigment-type switching puzzle. Pigment Cell Melanoma Res., 23, 4–6. Sturm, R.A. (2002) Skin colour and skin cancer – MC1R, the genetic link. Melanoma Res., 12, 405–416. Sturm, R.A. (2009) Molecular genetics of human pigmentation diversity. Hum. Mol. Genet., 18, R9–17. Sturm, R.A., Duffy, D.L., Box, N.F., Newton, R.A., Shepherd, A.G., Chen, W., Marks, L.H., Leonard, J.H., and Martin, N.G. (2003) Genetic association and cellular function of MC1R variant alleles in human pigmentation. Ann. NY Acad. Sci., 994, 348–358. Sturm, R.A., Duffy, D.L., Box, N.F., Chen, W., Smit, D.J., Brown, D.L., Stow, J.L., Leonard, J.H., and Martin, N.G. (2003) The role of melanocortin-1 receptor polymorphism in skin cancer

339

340

11 Genetics of Melanosome Structure and Function

36

37

38

39

40

41

42

43

44

45

46

47

risk phenotypes. Pigment Cell Res., 16, 266–272. Norton, H.L., Kittles, R.A., Parra, E., McKeigue, P., Mao, X., Cheng, K., Canﬁeld, V.A., Bradley, D.G., McEvoy, B., and Shriver, M.D. (2007) Genetic evidence for the convergent evolution of light skin in Europeans and East Asians. Mol. Biol. Evol., 24, 710–722. McEvoy, B., Beleza, S., and Shriver, M.D. (2006) The genetic architecture of normal variation in human pigmentation: an evolutionary perspective and model. Hum. Mol. Genet., 15, R176–R181. Tobin, D.J. (2008) Human hair pigmentation – biological aspects. Int. J. Cosmet. Sci., 30, 233–257. Sturm, R.A. and Larsson, M. (2009) Genetics of human iris colour and patterns. Pigment Cell Melanoma Res., 22, 544–562. Orlow, S.J. and Brilliant, M.H. (1999) The pink-eyed dilution locus controls the biogenesis of melanosomes and levels of melanosomal proteins in the eye. Exp. Eye Res., 68, 147–154. Yamaguchi, Y., Morita, A., Maeda, A., and Hearing, V.J. (2009) Regulation of skin pigmentation and thickness by dickkopf 1 (DKK1). J. Invest. Dermatol. Symp. Proc., 14, 73–75. Choi, W., Wolber, R., Gerwat, W., Mann, T., Batzer, J., Smuda, C., Liu, H., Kolbe, L., and Hearing, V.J. (2010) The ﬁbroblast-derived paracrine factor neuregulin-1 has a novel role in regulating the constitutive color and melanocyte function in human skin. J. Cell Sci., 123, 3102–3111. Brenner, M. and Hearing, V.J. (2008) Modifying skin pigmentation – approaches through intrinsic biochemistry and exogenous agents. Drug Discov. Today Dis. Mech., 5, e189–e199. Jimbow, K. (1997) Tuberous sclerosis and guttate leukodermas. Semin. Cutan. Med. Surg., 16, 30–35. Ortonne, J.P. (1990) Pigmentary changes of the ageing skin. Br. J. Dermatol., 122 (Suppl. 35), 21–28. Boissy, R.E. and Manga, P. (2004) On the etiology of contact/occupational vitiligo. Pigment Cell Res., 17, 208–214. Spritz, R.A. (2007) The genetics of generalized vitiligo and associated

48

49

50

51

52

53

54

55

56

57

58

autoimmune diseases. Pigment Cell Res., 20, 271–278. Spritz, R.A. (2008) The genetics of generalized vitiligo. Curr. Dir. Autoimmun., 10, 244–257. Spritz, R.A. (2010) Shared genetic relationships underlying generalized vitiligo and autoimmune thyroid disease. Thyroid, 20, 745–754. Brenner, M., Coelho, S.G., Beer, J.Z., Miller, S.A., Wolber, R., Smuda, C., and Hearing, V.J. (2009) Long-lasting molecular changes in human skin after repetitive in situ UV irradiation. J. Invest. Dermatol., 129, 1002–1011. Choi, W., Miyamura, Y., Wolber, R., Smuda, C., Reinhold, W., Liu, H., Kolbe, L., and Hearing, V.J. (2010) Regulation of human skin pigmentation in situ by repetitive UV exposure – molecular characterization of responses to UVA and/or UVB. J. Invest. Dermatol., 130, 1685–1696. Dell’Angelica, E.C., Mullins, C., Caplan, S., and Bonifacino, J.S. (2000) Lysosomerelated organelles. FASEB J., 14, 1265–1278. Huizing, M., Anikster, Y., and Gahl, W.A. (2000) Hermansky–Pudlak syndrome and related disorders of organelle formation. Trafﬁc, 1, 823–835. Huizing, M., Boissy, R.E., and Gahl, W.A. (2002) Hermansky–Pudlak syndrome: vesicle formation from yeast to man. Pigment Cell Res., 15, 405–419. Li, W., Rusiniak, M.E., Chintala, S., Gautam, R., Novak, E.K., and Swank, R.T. (2004) Murine Hermansky–Pudlak syndrome genes: regulators of lysosomerelated organelles. Bioessays, 26, 616–628. Wei, M.L. (2006) Hermansky–Pudlak syndrome: a disease of protein trafﬁcking and organelle function. Pigment Cell Res., 19, 19–42. Dessinioti, C., Stratigos, A.J., Rigopoulos, D., and Katsambas, A.D. (2009) A review of genetic disorders of hypopigmentation: lessons learned from the biology of melanocytes. Exp. Dermatol., 18, 741–749. Marks, M.S., Theos, A.C., and Raposo, G. (2003) Melanosomes and MHC class II antigen-processing compartments: a tinted view of intracellular trafﬁcking and immunity. Immunol. Res., 27, 409–425.

References 59 Dell’Angelica, E.C. (2003) Melanosome

60

61

62

63

64

65

66

67

68

69

70

71

72

biogenesis: shedding light on the origin of an obscure organelle. Trends Cell Biol., 13, 503–506. Setaluri, V. (2003) The melanosome: dark pigment granule shines bright light on vesicle biogenesis and more. J. Invest. Dermatol., 121, 650–660. Hearing, V.J. (2005) Biogenesis of pigment granules: a sensitive way to regulate melanocyte function. J. Dermatol. Sci., 37, 3–14. Raposo, G. and Marks, M.S. (2007) Melanosomes – dark organelles enlighten endosomal membrane transport. Nat. Rev. Mol. Cell Biol., 8, 786–797. Park, H.Y., Kosmadaki, M., Yaar, M., and Gilchrest, B.A. (2009) Cellular mechanisms regulating human melanogenesis. Cell Mol. Life Sci., 66, 1493–1506. Yamaguchi, Y. and Hearing, V.J. (2009) Physiological factors that regulate skin pigmentation. BioFactors, 35, 193–199. Wu, X. and Hammer, J.A., III (2000) Making sense of melanosome dynamics in mouse melanocytes. Pigment Cell Res., 13, 241–247. Westbroek, W., Lambert, J.M., and Naeyaert, J.M. (2001) The dilute locus and Griscelli syndrome: gateways towards a better understanding of melanosome transport. Pigment Cell Res., 14, 320–327. Seabra, M.C., Mules, E.H., and Hume, A.N. (2002) Rab GTPases, intracellular trafﬁc and disease. Trends Mol. Med., 8, 23–30. Langford, G.M. (2002) Myosin-V, a versatile motor for short-range vesicle transport. Trafﬁc, 3, 859–865. Maniak, M. (2003) Organelle transport: a park-and-ride system for melanosomes. Curr. Biol., 13, R917–R919. Futter, C.E. (2006) The molecular regulation of organelle transport in mammalian retinal pigment epithelial cells. Pigment Cell Res., 19, 104–111. Coudrier, E. (2007) Myosins in melanocytes: to move or not to move? Pigment Cell Res., 20, 153–160. Aspengren, S., Hedberg, D., Skold, H.N., and Wallin, M. (2009) New insights into melanosome transport in vertebrate pigment cells. Int. Rev. Cell Mol. Biol., 272, 245–302.

73 Van, G.M., Dynoodt, P., and Lambert, J.

74

75

76

77

78

79

80

81

82

83

84

(2009) Griscelli syndrome: a model system to study vesicular trafﬁcking. Pigment Cell Melanoma Res., 22, 268–282. Cardinalli, G., Ceccarelli, S., Kovacs, D., Aspite, N., Lotti, L.V., Torrisi, M.R., and Picardo, M. (2005) Keratinocyte growth factor promotes melanosome transfer to keratinocytes. J. Invest. Dermatol., 125, 1190–1199. Cardinali, G., Bolasco, G., Aspite, N., Lucarini, G., Lotti, L.V., Torrisi, M.R., and Picardo, M. (2008) Melanosome transfer promoted by keratinocyte growth factor in light and dark skin-derived keratinocytes. J. Invest. Dermatol., 2008, 558–567. Boissy, R.E. (2003) Melanosome transfer to and translocation in the keratinocyte. Exp. Dermatol., 12, 5–12. Van Den Bossche, K., Naeyaert, J.M., and Lambert, J. (2006) The quest for the mechanism of melanin transfer. Trafﬁc, 7, 769–778. Tadokoro, T., Yamaguchi, Y., Batzer, J., Coelho, S.G., Zmudzka, B.Z., Miller, S.A., Wolber, R., Beer, J.Z., and Hearing, V.J. (2005) Mechanisms of skin tanning in different racial/ethnic groups in response to ultraviolet radiation. J. Invest. Dermatol., 124, 1326–1332. Levine, N. (1993) Pigmentation and Pigmentary Disorders, CRC Press, Boca Raton, FL. Prota, G. (1992) Melanins and Melanogenesis, Academic Press, New York. Zeise, L., Chedekel, M.R., and Fitzpatrick, T.B. (1995) Melanin: Its Role in Human Photoprotection, Valdenmar, Overland Park, KS. Fitzpatrick, T.B., Szabo, G., Seiji, M., and Quevedo, W.C., Jr (1976) Biology of the melanin pigmentary system, in Dermatology in General Medicine (eds T.B. Fitzpatrick, A.Z. Eisen, K. Wolff, I.M. Freedberg, and K.F. Austen), McGrawHill, New York, pp. 131–163. Brenner, M. and Hearing, V.J. (2009) What are melanocytes really doing all day long..? Exp. Dermatol., 18, 799–819. Tachibana, M. (1999) Sound needs sound melanocytes to be heard. Pigment Cell Res., 12, 344–354.

341

wwwwwww

343

12 Physiological and Pathological Functions of Melanosomes Jan Borovanský and Patrick A. Riley

12.1 Tissue Concentration of Melanosomes

The behavior of pigmented tissues is in some important respects speciﬁc due to the presence of melanosomes, and reﬂects the chemical composition and tissue concentration and degree of dispersion of these specialized organelles [1]. There have been many reports, using spectrometric and chemical analyses, detailing the tissue melanin concentration and the distinction between the proportion of eumelanin and pheomelanin that have been reviewed by Ito and Wakamatsu [2]. Reﬂectance spectroscopy, used to measure changes in pigmentation, ascertains relative alterations in melanin level rather than the melanosome concentration [3, 4]. Only recently, Nielsen et al. [5], using an accurate discrete ordinate radiation transfer model for the coupled air–tissue system, known as the s.c.CATDISORT model, in conjunction with a classical inversion scheme based on Bayesian optimal estimation theory for retrieval of parameters, proved that from reﬂectance spectra it is feasible to calculate the melanosome concentration in the lower and upper epidermis. Some information on the quantity of melanosomes in individual cells can be obtained by three-dimensional imaging of mammalian cells with ion-abrasion scanning electron microscopy [6]; however, owing to the impracticality of their direct quantiﬁcation on a large scale, there has hitherto been only one attempt to assess the tissue concentration of melanosomes. Borovanský et al. [7] developed an indirect approach by combining the estimation of the melanin concentration in freeze-dried tissues and that in melanosomes isolated from them. From these data the concentrations of melanosomes found in various pigmented tissues were calculated (Figure 12.1). The estimated melanosome concentrations were found to be consistent with the depth of pigmentation of the analyzed tissues and also with their capacity to perform various functions ascribed mainly to the presence of melanin.

Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

344

12 Physiological and Pathological Functions of Melanosomes

Tissue concentration of melanosomes in freeze-dried tissue [7–10]. dHH, dark human hair; bHH, black human hair; bDH, black dog hair; BC, bovine choroid; BHM, Bomirski hamster melanoma; HP, Harding-Passey mouse melanoma; B16, B16 mouse melanoma; HM, human melanoma; EQ, horse melanoma; mPM, minipig melanoma.

Figure 12.1

12.2 Melanosome Properties and Functions Are Determined by Their Chemical Composition

The ﬁrst data on the chemical composition of melanosomes were not obtained until the problem of separation of intact and homogenous samples of melanosomes had been solved in 1960s [11, 12]. The early studies [8, 9], based on acid hydrolysis of melanosomes, gravimetric determination of melanin, and sum of amino acids recovered by the amino acid analysis, revealed that the main melanosomal constituents are melanin and protein (Figure 12.2), and indicated the presence of other components, such as lipid, carbohydrates, and inorganic compounds, the presence of which was conﬁrmed later. Ganglioside proﬁles of melanosomes isolated from B16 mouse melanoma and Bomirski Syrian hamster melanoma were compared with those of melanoma cells. The isolated melanosomes contained GM3 and GD3 as their major ganglioside components, extending the group of antigenic determinants shared by melanosomes and the cell surface of melanoma cells (see also Section 12.8) [13]. Comparison of lipids between melanosomes isolated from Syrian hamster eyes and Bomirski Syrian hamster melanoma showed that tumor melanosomes contained 8–10% of total lipids, half of which were phospholipids, unlike normal melanosomes that contain 3–4% of total lipids, among which no phospholipids were detected [14].

12.2 Melanosome Properties and Functions Are Determined by Their Chemical Composition

HP

B16

HU

RP

S91

EQ

OX

Chemical composition of melanosomes [8–10]. Black segments = melanin, gray segments = protein, white segments = other substances. Top row: melanosomes from mouse melanomas (HP, Harding-Passey; B16; S91, Cloudman S91; ST, Stanford). Middle row: tumor melanosomes (HU, human melanoma; EQ, horse melanoma; MA, Bomirski Syrian hamster melanoma). Bottom row: melanosomes of

Figure 12.2

ST

MA

SQ

SE

nontumor origin (RP, retinal pigment epithelium of chicken embryos; OX, bovine choroid; SQ, ink granules from squid (Lolio pealii); SE, ink granules of Sepia ofﬁcinalis). All the samples were prepared by the method of Seiji et al. [11] and the purity of the samples was checked by electron microscopy. Jimbow et al. [12] obtained exactly the same results with B16 melanoma melanosomes and Sepia melanin granules.

As for the protein moiety of melanosomes, research activities naturally at ﬁrst focused on proteins with enzyme activities, primarily tyrosinase and its related proteins (see Chapter 4). In parallel, many acid hydrolases were detected as common melanosomal constituents [15], summarized by Borovanský and Elleder [16], whereas structural proteins were characterized later (see Chapter 9). Nowadays the secondary structure of many melanosomal proteins is known and their genes have been cloned (see Chapter 11), and protein trafﬁcking and targeting to melanosomes has been described in detail (see Chapter 9). The dawn of proteomic techniques has increased by several orders of magnitude our possibilities to detect and characterize residual molecules, molecules in transit, and molecules that transiently interact with the organelle to carry out distinct functions [17]. Proteomic analysis of early melanosomes from MNT-1 cells made by Basrur et al. [18] identiﬁed all ﬁve melanosome-speciﬁc proteins (tyrosinase, tyrosinase-related protein 1 (TRP-1), TRP-2 (also known as dopachrome tautomerase (Dct)), MART-1, and OA1) together with 56 proteins shared with other cell organelles, and revealed the presence of six novel melanosomal proteins. Proteomic study of melanosomes isolated from pigmented MNT-1 and nonpigmented SK-Mel-28 human melanoma cells [19] at various developmental stages identiﬁed around 1500 proteins in melanosomes of all stages, with around 600 at any given stage. Approximately 100 proteins shared by melanosomes from pigmented and

345

346

12 Physiological and Pathological Functions of Melanosomes

nonpigmented cells deﬁned the essential melanosome proteome. Comparative bioinformatics analyses and proﬁling of lysosome-related organelles proteomes provides a basis for further hypothesis formulation and experimental validation of organelle proteins and their functional roles [20].

12.3 Functional Microanatomy of the Melanosome

In vertebrate melanosomes (schematically depicted in Figure 12.3) the pigment is arranged along a protein matrix in a succession of small quasispherical granules approximately 30 nm in diameter. There is some evidence that the granules exhibit a degree of internal heterogeneity with a core of pheomelanin and a coating of eumelanin (see Section 6.3.4). This may be ascribed to the bias of the competing rate constants for dopaquinone reactions that favors reaction with thiols leading to pheomelanin precursors over cyclization leading to eumelanin precursors. However, since this occurs successively at different intramelanosomal locations such an explanation requires spatially organized concentrations of thiol groups. Contrary to earlier views of the structure of melanins there does not seem to be any large-scale polymerization of melanogenic precursors, and the current consensus is that the basic melanin units are generally oligomers of between three and six subunits [21] which, in the case of eumelanin, consist of 5,6-dihydroxyindole (DHI) or DHI-2-carboxylic acid (DHICA) monomers and their respective quinones [22]. The relative stability of these quinones is of some interest and may depend on delocalization of electrons within the oligomers. Within a 30-nm diameter granule there is potential space for about 2.5 × 106 oligomeric melanin units and the cohesion of these units seems to depend on their organization on the protein matrix. The arrangement of melanin units has the physical characteristics of a

Schematic outline of the organization of melanosomes bounded by a lipid membrane and containing a protein matrix along which melanin particles are arranged (not to scale).

Figure 12.3

12.3 Functional Microanatomy of the Melanosome

disorderly array [23] and the processes involved in the aggregation of oligomers are not known. However, a number of possibilities have been suggested and it is possible that several are involved. These include: i) ii) iii) iv) v) vi)

Reaction of nascent quinones with cysteine residues of a matrix protein. Reaction of nascent quinones with free amino groups of a matrix protein. Copolymerization of oligomers with oxidized tyrosyl residues of protein. Hydrogen bonding. Hydrophobic interactions. Formation of ionic or charge-transfer complexes.

Protein binding through quinone reactivity with free thiols has been proposed [24] (Figure 12.4) and this may be a means of attachment of melanin units to the underlying structural matrix. Amino acid analysis of melanosomal protein indicates that there are signiﬁcant numbers of cysteine residues. In a similar fashion, reaction with free amino groups, such as lysine residues (Figure 12.5), may anchor oligomeric melanin units to matrix proteins [8, 9, 26, 27].

Figure 12.4 Electrophilic addition of dopaquinone to a cysteine residue in a protein chain. (Based on the scheme in Chedekel et al. [25].)

Figure 12.5 Addition of dopaquinone to the terminal amino group of a lysine residue in a protein chain.

347

348

12 Physiological and Pathological Functions of Melanosomes

Another possibility for attachment to matrix proteins is through copolymerization with oxidized tyrosyl residues (Figure 12.6) [28, 29]. In addition to covalent binding to protein there are other possible modes of adhesion including hydrogen bonding through the hydroxyl or carbonyl groups of DHI and DHICA. Hydrogen bonds could form between melanin oligomers depending on the availability of quinone groups, and also between melanin and suitable hydrogen bond donors or acceptors of proteins, including the hydrogenbonding functions of the peptide chain (Figure 12.7). There is evidence that oligomeric units may associate to form stacks [21] and similar interactions with hydrophobic regions of proteins could impose some structural cohesion. Finally, the existence of many ionizable groups on the melanin oligomers suggests the possibility that ionic complexes could contribute signiﬁcantly to the three-dimensional structure of the granular aggregates. It is well recognized that melanin exhibits properties of a cation-exchange material and strongly binds polyvalent metals. This property would permit complexes to form between the

Oxidation of a tyrosyl residue by tyrosinase with subsequent incorporation of the dihydroxyindole moiety into melanin.

Figure 12.6

12.3 Functional Microanatomy of the Melanosome

Figure 12.7 Possible hydrogen bonding between indolic oligomers and protein.

associated oligomers. It has been shown that metal addition inﬂuences the structure of synthetic melanin [30], which suggests that the aggregation patterns of melanin oligomers may be sensitive to the type and concentration of metal ligands. Such complexes could involve ionized groups on proteins, or metals ligated to proteins, thus imposing some structural coherence and the possibility of chargetransfer complexes may have signiﬁcance to the spectral properties of melanin. Clearly there are multiple possibilities for interaction of melanin oligomers with protein, and the signiﬁcance of cysteine, arginine, histidine, tyrosine, and serine in melanin binding is supported by the effects of methylation of these amino acids on the increased solubility of melanins and alterations in spectral properties [31]. The possible role of hydrogen bonds or ionic bonds in determining the structure of melanin is suggested by the sensitivity of the pigment to hydration. Degradation of melanin is accelerated by hydration and the physical properties are altered [32]. Consideration of the interactions between melanin and the protein moiety of melanosomes revives the dispute concerning the factors responsible for the shape of melanosomes [33]. Some Japanese authors [34, 35] have proposed that the ultrastructure of the melanosome is a secondary consequence of the type of pigment generated and that ovoid melanosomes with lamellar ultrastructure result from eumelanin generation, whereas pheomelanin synthesis leads to spherical melanosomes with granular ultrastructure. However, Harding-Passey melanoma melanosomes, which characteristically contain eumelanin [36], exhibit granular ultrastructure, and the occurrence of lamellar melanosomes in red hair [37] do not support the proposed relationship between melanosome ultrastructure and the type of pigment produced. From the very beginning we have adhered [38, 39] to the suggestion of Moyer [40] that the major determinant of the ultrastructure of the melanosome is not the type of pigment produced, but the nature of the matrix protein(s). Melanin produced in the absence of melanosomal matrix proteins in

349

350

12 Physiological and Pathological Functions of Melanosomes

tyrosinase-transfected ﬁbroblasts is deposited irregularly in lysosomes [41] and clearly indicates the decisive role of proteins in regulating the shape of melanincontaining organelles. There is evidence that the melanosomal protein, Pmel17 (see Section 9.3), acts as a framework for melanin deposition. It seems logical, that the ontogenetically older melanosomal moiety (i.e., proteins) will inﬂuence the structure of the ontogenetically younger part of the melanosome (i.e., melanin) and it is astonishing that the idea that the type of melanin shapes the melanosome has persisted in recent literature.

12.4 Melanosomes as Centers of Free Radical Activity 12.4.1 Free Radical Nature of Melanins

Under normal circumstances in melanocytes melanin is deposited solely in melanosomes. Both basic types of authentic melanins – eumelanins and pheomelanins – belong to the class of stable free radicals. The free radical content is lower in eumelanin than in pheomelanin. The free radical content of melanins is typically around 2 × 1018 spins/G for an aqueous suspension at neutral pH [42]. In physiological conditions the melanin radicals in melanosomes are not harmful as the radicals are incorporated in an insoluble biopolymer and, therefore, of limited mobility. The radical nature, especially of eumelanosomes, has been characterized as cytoprotective [43] with at least three mechanisms involved [44]: i) As melanins in melanosomes are redox biopolymers containing large amounts of oxidizing (o-quinone) and reducing (o-hydroquinone groups) [42, 44], they can act as sinks for diffusible radical species: a) by electron donation, for example, reaction with superoxide anion radical: reduced melanin + O2 i − + 2H+ → H2O2 + melanin radical b)

by electron capture, for example: oxidized melanin + O2 i − → O2 + melanin radical

ii) At locations exposed to light (such as the skin and the eye) melanosomal melanin, by converting light energy into vibrational energy of molecules (for details, see Section 12.5.1), prevents its absorption by other chromophores whose excited states would lead to the generation of singlet oxygen [45]. This mechanism is often called the ﬁlter effect [46]. iii)

Melanosomes by binding redox active metal ions (see Section 12.6) may alter their redox potential and/or their accessibility, by these means shifting oxidative stress to the melanosomal compartment and thus sparing more critical cellular targets [44, 47]. The cytoprotective function of eumelanin has been tested in practice. Melanin-covered silica nanoparticles were successfully

12.4 Melanosomes as Centers of Free Radical Activity

tested for radioprotection of bone marrow during radiation therapy of cancer in an animal model [48]. Melanosomes are also able to exert cytotoxic effects and this applies especially to those organelles containing pheomelanin. Pure pheomelanin is not observed in nature and rarely makes up more than 25% of the total melanin present [49]. Measurements of the surface photoionization threshold of intact melanosomes have revealed a lower photoionization potential for melanosomes containing pheomelanin in comparison with eumelanosomes [49] and the optical absorption coefﬁcient of intact melanosomes decreases with increasing pheomelanin content [49]. The recently proposed casing model concept of mixed melanogenesis in melanosomes (see Chapter 6), with a pheomelanin center coated by eumelanin, makes the overall free radical reactivity more complex. The common belief that eumelanin acts as a cytoprotective and photoprotective antioxidant while pheomelanin has a phototoxic pro-oxidant action has persisted, but the actual behavior of melanin requires a more detailed evaluation. The majority of melanin free radicals (e.g., semiquinones) exist in equilibrium with nonradical polymer units such as the corresponding quinones and catechols, and various conditions, such as temperature, pH, or illumination, determine the comproportionation equilibrium and affect the free radical concentration. If the energy input is too high, so as to exceed the capacity of the pigment to detoxify radicals, instead of having a cytoprotective action, melanin may participate in radical production with possible cytotoxic consequences [44]. However, the biological importance of melanin as a “photoprotective” agent may reside in such an effect in that it ensures the destruction of cells exposed to radiation of sufﬁcient energy to cause deleterious mutations [50]. With respect to the role of melanins and melanosomes in photoprotection, the action involving light has received most attention (summarized by Borovanský [44] and Sarna and Swartz [51]). Irradiation of melanins with either visible or UV light may produce an excited state (M(•)n*), which in turn yields melanin with increased free radical activity (M(•)n + x). Irradiation of melanin has been reported to increase the free radical content which returns to the ground level following the discontinuation of the irradiation. Since melanin can undergo facile single-electron oxidation and reduction, it is conceivable that excited-state melanin reacts with molecular oxygen to yield superoxide O2•− [52, 53] and the oxidized form of melanin (M(•)nox). Such a process can be reversed in the presence of reducing compounds present in the melanocyte [54]. Alternatively the interaction of excited-state melanin with O2 may result in the transfer of excitation energy to oxygen yielding singlet oxygen 1 O2. As melanin has been shown to quench excited states of other molecules, such as photosensitizers (S), the production of singlet oxygen can be observed also in sensitized photolysis. Mechanisms of free radical generation during photoreactions of melanin are summarized as:

•

Direct photoreactions of melanin: M(i)n + hν → M(i)n * → M(i)n + x

351

352

12 Physiological and Pathological Functions of Melanosomes

M(i)n * + O2 → M(i)n ox + O2i − M(i)n + x + O2 → M(i)n ox + O2 i −

•

Sensitized photoreactions: S + hν → S* S* + M(i)n → S + M(i)n + x S* + O2 → S + 1O2

Irradiation of melanins in vitro in the presence of air was shown to promote oxygen consumption and production of superoxide and hydrogen peroxide [46, 52]. Hydrogen peroxide presents a potential threat due to the possibility of hydroxyl radical generation via the metal-catalyzed Haber–Weiss reaction; under some circumstances hydroxyl radicals can be produced directly by UV irradiation of melanins [55]. Production of superoxide is much higher in melanoma cells compared to melanocytes [56]. Superoxide ions and hydrogen peroxide are involved in photodegradation of melanosomes and melanin [16, 57]. Menon et al. [58] demonstrated that pheomelanin produces large amounts of reactive oxygen species (ROS) and melanin radicals when irradiated with UV/visible light. This is manifested by cytotoxic effects, including lipid peroxidation, and inﬂammatory reaction of the skin [46]. 12.4.2 Radicals and Reactive Species Associated with Melanogenesis

Examination of the Raper–Mason scheme (see Chapters 3, 4 and 6) discloses that melanogenesis is a potential hazard for melanocytes that are endangered by potentially cytotoxic intermediates – quinones and semiquinones – in two main ways. Quinones, being highly electrophilic, can bind covalently with proteins through nucleophilic thiol groups [59] (see also Sections 12.4.1 and 12.4.3). If this were to occur extramelanosomally, inhibition of a number of enzymes with important cellular functions, such as DNA polymerase, would ensue [60, 61]. Soluble reaction products of tyrosinase-catalyzed oxidation have been shown to bind covalently to DNA [60]. Everybody who has experience with DNA isolation from pigment cells knows that isolated DNA is slightly pigmented! Semiquinone species (dopa semiquinone, cyclodopa semiquinone, and indole-2-carboxylic acid semiquinone) can be generated during tyrosinase-catalyzed melanogenesis by redox equilibration between quinones, and catechols (Eq. 12.1) [62], and may initiate lipid peroxidation in melanocytes and destroy them [63] (Figure 12.8). One-electron oxidations of melanin precursors by peroxidase/H2O2 (Eq. 12.2) and during autoxidation (Eq. 12.3) can also generate semiquinones (reviewed by Sealy [42]):

12.4 Melanosomes as Centers of Free Radical Activity

Figure 12.8 Schematic outline of the initiation of lipid peroxidation by semiquinone radical.

catechol + o-quinone ↔ 2 semiquinones + 2H+

(12.1)

2 catechols + H2O2 → 2 semiquinones + 2H2O + 2H+

(12.2)

−

catechol + O2 i→ semiquinone + O2 + 2H

+

(12.3)

Under normal circumstances the potentially dangerous process of melanogenesis is strictly compartmentalized within melanosomes where the conversion of diffusible reactive intermediates into the insoluble melanin biopolymer takes place and where nonspeciﬁc binding of melanin precursors to proteins provides an additional means of scavenging reactive intermediates [44, 64]. In melanoma cells the occurrence of aberrant melanosomes is typical [65] and includes defects in, or even complete absence of, the limiting membrane [64, 66, 67] (Figure 12.9). Although the toxicity of melanin precursors [68] and membrane defects of tumor melanosomes have long been known, it was only in the early 1990s that these facts were put together and the concept of leaky melanosomes with cytotoxic consequences was proposed [33, 44, 61, 64] (Figure 12.10). Offner et al. [69] have demonstrated in vitro that human melanoma cells (unlike other tumor cells) were able to damage endothelial cells by an oxygen radical-dependent mechanism and suggested that melanogenesis-derived radical species might be involved in the metastatic propensity of melanoma cells. Under physiological conditions leakage of toxic melanin precursors is limited because the permeability of melanosomal membrane is low [70]. In addition, pigment cells possess physiological scavenging mechanisms to cope with toxic species that might gain access to the cytosol. Quinone species may be scavenged by reaction with cysteine [59] or glutathione [71] forming 5-S-cysteinyldopa and

353

354

12 Physiological and Pathological Functions of Melanosomes

Electron microscope appearance of aberrant melanosomes in human malignant melanoma showing serious defects or even a complete absence of the limiting membrane.

Figure 12.9

a)

b)

Figure 12.10 Scheme of intracellular distribution of metabolites in normal melanocytes and in melanoma cells with defective melanosomes. The reactants are abbreviated as: S, substrate (e.g., tyrosine); Q, quinone; M, melanin biopolymer; RSH, nucleophilic thiol species (e.g., cysteine or glutathione); RSQH, addition product (cysteinydopa or glutathionyldopa). (a) Normal situation in the melanocyte: the melanosomal membrane prevents escape of quinones into the cytosol and any leaked quinones are easily scavenged by cytosolic

scavengers and excreted in the urine. (b) Pathological situation in melanoma cells: in the case of melanosomes with membrane defects access to tyrosinase substrates is facilitated, as is the diffusion of reactive quinones into the cytosol. Removal of quinones by reaction with scavenging molecules depletes the cell of antioxidant capacity and generates a large amount of excretion products. When the capacity of scavenging mechanisms is overwhelmed, the reactivity of unscavenged toxic species is responsible for cytotoxic phenomena.

12.4 Melanosomes as Centers of Free Radical Activity

glutathionyldopa, which can be converted to 5-S-cysteinyldopa, which can be excreted in the urine. It is important to realize that there are two pools of 5-Scysteinyldopa in melanocytes: a melanosomal pool (used to synthesize pheomelanins) and a cytoplasmic pool (a manifestation of the detoxiﬁcation process). The potentially toxic dihydroxy derivatives of melanogenesis, such as DHI [72] and DHICA, can be detoxiﬁed by catechol-O-methyltransferase (COMT) [73]. In melanoma cells containing melanosomes with membrane defects, the leakage of toxic melanin precursors is high and the physiological scavenging mechanisms work at full capacity, which is reﬂected by increased excretion of melanogens [74, 75], and, as soon as their capacity is overwhelmed, pathological mechanisms ensue, including extramelanosomal melanin deposition, quinone binding to -SH groups with enzyme inactivation, DNA damage (genotoxicity), lipid peroxidation, and other phenomena (summarized by Borovanský [44, 64]). The degree of permeability of the melanosomal limiting membranes, resulting from both structural defects and damage by intramelanosomal free radical reactions, may determine the extent of cytotoxic phenomena observed in melanoma cells and may also be one of the factors determining the extent of melanogenuria. Participation of autocytotoxic phenomena due to melanosome membrane defects might be also be a factor in the phenomenon of spontaneous melanoma regression [69]. In this context, the possibility of amplifying the generation of toxic melanogenic intermediates has long been viewed as a basis for a rational approach to melanoma therapy (see Section 12.4.4). 12.4.3 Possible Role of Protein-Bound Dopa

Analyses of melanosomal hydrolysates regularly reveal the presence of 3,4-dihydroxyphenylalanine (dopa) [8, 76]. In mammalian cells protein-bound dopa can be generated by both controlled enzymatic pathways and by uncontrolled radical reactions [77]. Moreover, although dopa-tRNA has not been described, free dopa may be incorporated into protein by protein synthesis [78]. Mushroom tyrosinase can catalyze hydroxylation of tyrosyl residues in proteins to dopa and subsequent oxidation to dopaquinone residues [79]. Protein-bound dopa and protein hydroperoxides are among the major long-lived redox active products formed during free radical attack on proteins [80], and such a milieu inside melanosomes is to be expected. Protein-bound dopa can redox cycle between catechol and quinone forms, and bind transition metals [81]. Protein-bound dopa, behaving as a redox-active product, is capable of functioning as both a pro- and antioxidant [81]. Unlike free dopa, the levels of protein-bound dopa can change 5- to 10-fold during oxidative damage in vivo, which is an appropriate property for a signaling molecule. Nelson et al. [80] have suggested a mechanism by which protein-bound dopa might trigger antioxidant defense via nuclear factor NF-κB and other transcription factors. With the exception of a study investigating the release of radioactivity from isolated melanosomes metabolically labeled with [2-14C]l-dopa [82], no attempt (using pigment cells) has been made to verify the hypothesis that the

355

356

12 Physiological and Pathological Functions of Melanosomes

generation of protein-bound dopa can trigger an enhancement of the cellular antioxidant defense, thus minimizing the level of oxidative damage [83] or the contrary view that protein-bound dopa is able to promote further radical-generating events such as oxidative DNA damage [84, 85]. 12.4.4 Melanosomes as a Therapeutic Target

The reactivity of o-quinones has been noted in Chapter 3 and it is likely that the evolutionary advantage of conﬁning melanogenesis to a specialized organelle follows from the havoc that could follow from permitting free diffusion of the quinone intermediates throughout the cytosol. The signiﬁcance of this was expressed by Hochstein and Cohen [68] in the following words: We have in mind the fact that tyrosine sequestered in melanotic tumours is converted to melanin through the intermediate formation of many polyphenolic and quinoid intermediates, for example, 3,4-dopa, 5,6-dihydroxyindole and their corresponding quinones. Such substances belong to a class of potentially cytotoxic agents. An extensive literature exists on their inhibitory action in a variety of microbial and mammalian systems. One may therefore view the melanotic cells as being continuously challenged by toxic agents derived from tyrosine during the process of melanin formation. The importance of containment of melanogenesis in the melanosomal compartment is clear and to some extent is achieved by the presence of a lipid membrane that forms a relatively impermeable barrier to the diffusion of small molecules, particularly if they are charged. In addition, there are several detoxifying mechanisms that are brought into play to minimize the damage that could result from their escape into the cytosol. These include quinone reductase, glutathione S-transferase and COMT, as reviewed by Smit et al. [84]. In addition, because of the redox reactivity and other properties of melanins, the containment of the melanin product of the oxidative reactions is also of importance. Given these reasons for the intracellular segregation of melanogenesis it is of considerable consequence that defective melanosomes with incomplete limiting membranes have been detected in melanomas [64], and these properties of melanosomes lend themselves to exploitation both in the diagnosis and treatment of melanoma [1]. Moreover, deﬁciencies of protective metabolic routes, such as those dependent on cellular glutathione levels [85], are thought to exist (see Chapter 13). These considerations suggest that melanogenesis could provide a possible means of a speciﬁc attack on melanoma cells [86]. In particular, given the usually raised level of tyrosinase activity in malignant melanocytes, targeted cytotoxicity could be exerted by leakage of quinones into the cytosol with consequent depletion of cellular thiols, damage to nucleic acids, or attack on proteins. Although such examples of cytotoxicity may be associated with normal melanogenic inter-

12.4 Melanosomes as Centers of Free Radical Activity

mediates, the reactivity of the quinones formed renders their diffusion range very limited and thus, even with defective containment by the melanosomal membrane, the concentration reaching the cytosol will be relatively small. Consequently, indirect methods of utilizing the melanogenic pathway have been adopted. The ﬁrst approach, which may be termed the “Achilles Heel” mechanism, involves the introduction of analog substrates for tyrosinase that are unable to undergo the intramolecular cyclization of dopaquinone, a process that occurs with a rate constant of 3.8 s−1 [87]. This class of noncyclizing compounds includes the depigmenting tyrosine analogs 4-hydroxyanisole and monobenzone, which are known to be oxidized by tyrosinase to give rise to the corresponding o-quinones [88, 89]. Since the corresponding o-quinones are unable to cyclize they are less likely to be incorporated into melanin oligomers and, as a result, may have a more extended diffusion range. Nevertheless, the in vivo dosage levels that might be achieved are unlikely to be in the range of quinone concentrations adequate for direct cytotoxicity [90, 91] and it is likely that the responses elicited are due to a haptenic effect due to quinone binding to protein with subsequent induction of an immune response. This is the most probable mechanism to account for the industrial depigmentation by monobenzone [92], and the experimental depigmentation produced by 4-hydroxyanisole and other phenols [93]. A similar mechanism may account for the depigmentation of vitiligo. Another group of tyrosinase-substrates, the phenolic thioether amines (N-acetyl4-S-cysteaminyl phenol and its acetyl derivative and N-propionyl derivatives), have been studied by Jimbow et al. [94] with promising cytotoxic effects in vitro. Further experiments were performed with N-propionyl-4-S-cysteaminylphenol (NPrCAP) attached to magnetite nanoparticles combining the cytotoxicity of oxidized NPrCAP with targeted intracellular hyperthermia upon exposure the nanoparticles to an alternating magnetic ﬁeld. Such treatment caused cytotoxic reactions as well as heat-shock responses, leading to elicitation of antitumor immune responses [95]. An alternative strategy involving the exploitation of melanosomes as targets in antimelanoma therapy has been based on prodrug activation by tyrosinase. This “Trojan Horse” approach has largely centered on the effect of dopaquinone cyclization on labilizing hydrolysable bonds, thereby releasing known cytotoxic agents. The basic mechanism outlined by Jordan et al. [96] employs an amide linkage to the side-chain of tyrosine or dopa that, as a result of the electronic movements associated with cyclization of the corresponding quinone, becomes vulnerable to hydrolytic cleavage, thus releasing any suitable agent attached through the amide (Figure 12.11). A number of encouraging studies have examined this approach [97–100], although there are problems associated with the efﬁciency of the release mechanism [101] and methods have been explored to increase the stabilization of the prodrug compounds. Some prodrugs are sensitive to enzymatic attack that renders them nonspeciﬁc and some agents are intrinsically unstable [102]. Progress has been made in devising protective groups [103] and alternative release mechanisms have been proposed [104]. However, thus far there have been no clinical studies to test the therapeutic efﬁcacy of this approach to melanoma treatment.

357

358

12 Physiological and Pathological Functions of Melanosomes

Figure 12.11 Melanocyte-directed enzyme prodrug therapy (MDEPT) based on labilization of amide bond. The scheme shows a drug bound through a hydrolysable link to the amino group of dopamine. On tyrosinase-catalyzed oxidation to the quinone

the amino nitrogen cyclizes and permits hydrolytic attack on the bond, releasing the drug. The lone-pair electrons on the nitrogen are shown and the curly arrows indicate the electron displacements.

12.5 Melanosomes as Energy Transducers

Melanin absorbs light over a wide range of spectral energies. The current model of the structure of melanin views it as a disorganized aggregate of oligomers principally, in the case of eumelanin, composed of indolic monomers. Melanin can not only absorb light, but also scatter it. The pattern of light scattering depends on the particle size. Melanosomes with a diameter of 300 nm mainly cause forward scattering of UV light in contrast to smaller particles such as melanin particles in keratinocytes (below 30 nm), which display a symmetrical scattering proﬁle [105]. Light scattering accounts for less than 6% of the total absorbance [106] and most of the light-absorbing properties of the melanin aggregates are due to electronic effects. The broad underlying spectral absorption is best explained if melanin is regarded as an assemblage of heterogeneous chromophores able to interact electronically, a feature of which is its bathochromicity (see Chapter 3). Fluorescent emission from melanin is extremely small [107] and more than 99.9% of the energy from absorbed photons is rapidly dissipated nonradiatively. This makes melanosomes important organelles for the facile absorption of radiant energy and the very rapid photodynamics of eumelanin in particular means that there is almost complete conversion of the energy in absorbed photons into heat – a process referred to as photon/phonon conversion.

12.5 Melanosomes as Energy Transducers

12.5.1 Photon/Phonon Conversion

Melanosomes can convert absorbed photons into phonons. A phonon is deﬁned as a unit of vibrational energy. Phonons and electrons are the two main types of elementary particles or excitations in solids. Whereas electrons are responsible for the electrical properties of materials, phonons determine such things as the speed of sound within a material and how much heat it takes to change its temperature. Since heat is relatively easily dissipated, such a property is generally protective because it reduces the amount of light energy available for photochemical reactions in the skin and in the eye, and in both these locations melanosomes represent the main absorbing components of the cell. Interestingly, the photon/phonon conversion property of melanosomes is utilized by poikilotherms as a means of raising their body temperature, and the close anatomical relationship between melanocytes and superﬁcial blood vessels is marked in these animals. A similar solar heating phenomenon may explain the function of the pigment cells present in the well-developed pecten in migratory birds, although photochemical energy generation has also been suggested [108]. The effect of absorbed light energy clearly depends on the rate of energy deposition. If the rate of energy deposition is faster than the rate of thermal diffusion (thermal conﬁnement), then the temperature of the exposed tissue will rise and, if a critical temperature is reached (typically about 10 °C above basal), then thermal damage occurs. If the light energy is deposited faster than mechanical relaxation can occur (stress conﬁnement), then a thermoelastic pressure wave is produced and tissue is disrupted by shear forces or by cavitation-nonlinear effects [109, 110]. Model experiments with aqueous suspension of porcine retinal pigment epithelium melanosomes revealed that short pulsed laser irradiation caused heating of melanosomes (to 150 °C) leading to surrounding liquid vaporization. The resulting microbubbles on the melanosome surface could be directly imaged by fast ﬂash-light photography [111]. Short-pulse laser irradiation in the nanosecond to picosecond time domain caused transient microbubble formation around melanosomes in the retinal pigment epithelium (RPE) cells [112]. For single 12-nm pulses the threshold for bubble detection was the same as the ED50 threshold for cell death [113]. The principle of photon/phonon conversion is clinically used in retinal diseases to selectively damage the RPE without affecting the neural retina, the photoreceptors, and the choroid [113, 114]. If the absorption of energy is high enough, even explosive vaporization of melanosomes in situ in skin can be brought out. It manifests as a visible whitening of the superﬁcial epidermal layer due to stratum corneum disruption [115]. The rapid formation and contraction of bubbles around melanosomes, described above, is not a unique feature of melanin-containing structures, but a general phenomenon. It can be similarly induced in functionalized nanoparticles by laser or electromagnetic waves. The rapid formation and contraction of bubbles around heated nanoparticles, associated with the propagation of pressure waves, could

359

360

12 Physiological and Pathological Functions of Melanosomes

bring about thermo-mechanical damage to the cell or to surrounding cells [116]. The strategy of inducing intracellular hyperthermia has become an interesting direction for future cancer therapy [117, 118]. The advantage of pigmented tumors is the fact that melanosomes are already present to permit this manipulation. 12.5.2 Photochemical Reactions

At relatively low energy levels (in the spectral range above about 300 nm) the small amount of energy not dissipated as heat is able to take part in two types of photochemical reaction:

•

One is the reversible electron transfer between reduced and oxidized subunits of melanin, which has the effect of altering the comproportionation equilibrium of quinone and hydroquinone units, and results in the generation of semiquinone radicals. Due to the numerous aromatic oligomers containing multiple π-electron systems, the generated Compton recoil electrons gradually lose energy while passing through the pigment, until their energy is sufﬁciently low to be trapped by stable free radicals present in melanin. Controlled dissipation of high-energy recoil electrons by melanin prevents secondary ionization and the generation of damaging free radical species [119].

•

In the presence of extrinsic electron donors or acceptors more complex reactions may take place, including the initiation of photochemical damage. Under aerobic conditions photo-oxidation can give rise to ROS, which are potentially cytotoxic to irradiated tissue.

Other photochemical reactions of melanin, such as photoionization and photohomolysis, are known to occur as the result of absorption of higher-energy photons in the spectral range 240–300 nm [120]. 12.5.3 Sound/Heat Conversion

Lyttkens et al. [121] observed that persons with more pigment in the inner ear coped better with noise compared to those with less-pigmented inner ears. They suggested an explanation based on the conversion of sound energy to an electronically excited state of melanin – an effect similar to the absorption of electromagnetic energy – with the subsequent dissipation of this energy as heat.

12.6 Melanosomes and Metal Ions

One of the important properties of eumelanins is binding of cations [122–124]. The ability of melanin to serve as a reservoir for metal ions – enabling storage,

12.6 Melanosomes and Metal Ions

release, and exchange – contributes to cellular homeostatic mechanisms; alternatively, the ability of melanin to strongly bind metals and sequester them within a segregated compartment (the melanosome) may act as a detoxiﬁcation mechanism for potentially toxic heavy metals, and possibly as an excretory pathway with signiﬁcance to hominid evolution [125, 126]. The presence of metals in melanosomes has been known from the very beginning of their investigation. Early studies (1955–1996) have been critically reviewed by Borovanský [127], and later studies discussed by Hong and Simon [128]. The metal content of ocular melanosomes has been reviewed by Eibl et al. [129] (see also Section 7.6.3). In order to understand the function of metals in living systems, cells, and their organelles, the chemical and biochemical bases of metal interactions with intracellular targets must be known [127]. Potts and Au [130] characterized the afﬁnity of 23 inorganic ions for melanin, and since then many studies have focused on melanin as a versatile cation-exchange biopolymer [131] controlling melanosomal ion content and cellular metabolism [1, 128, 129]. Analytical data derived from measurements on isolated melanosomes can be inﬂuenced by contamination during separation procedures or by a loss of metals during harsh isolation conditions. Hence, it is advantageous for the analytical data to be conﬁrmed by direct in situ determination of metals (e.g., by X-ray microanalysis [132] or by in vivo experiments with radioactive isotopes [133]). Excluding the question of accessibility, which differs according to the model system investigated, the amount of any speciﬁc cation bound to melanin will depend on a range of factors, including:

• • • •

The number of different types of potential binding site and the available numbers of each type of site. The afﬁnity of each type of binding site for the ion in question and the relative afﬁnities for competing ions. The combined afﬁnity of adjacent or coordinated sites. The relative concentrations of ions competing for the same sites.

Clearly these factors are dependent on the ions, the type of melanin, the type of binding site, and the physiological conditions of the assay. In the melanin moiety of melanosomes, various groups, including carboxyl, amino, phenolic, quinone, and semiquinone, are potentially able to serve as sites for metal binding [128, 134]. In certain deﬁned conditions Fe3+ is generally complexed to o-phenolic groups, while Ca2+ and Mg3+ are bound to carboxyl groups of DHICA-melanin [128, 134]; on this basis, differences in the Ca2+ and/or Mg2+ concentration have been used to estimate the relative amount of DHICA in melanin samples [129]. With the exception of zinc [135], non-melanin metalbinding sites have been less-frequently considered, although the presence of metals in catalytic centers of melanosomal enzymes, such as copper in tyrosinase and zinc in TRP-2, is well known. Three cations have received special attention: iron, zinc, and calcium. Melanin has a marked afﬁnity for iron and Liu et al. [136] described a greater than 400-fold increase in iron concentration in Sepia granules on incubation with an aqueous solution of FeCl3. There is some evidence that divalent iron is more

361

362

12 Physiological and Pathological Functions of Melanosomes

strongly bound [128]. This sequestration of Fe2+ ions may be important because of their possible involvement in free radical reactions, but the bound iron is probably still redox-active and able to take part in the Fenton reaction [137]. Nevertheless, the intramelanosomal generation of ROS may be viewed as less hazardous to the cell since H2O2 damage to melanin, hydroxylation of proteins, and lipid peroxidation would be to some extent contained. Melanosomes are unusually rich in zinc and, although the analysis of the afﬁnity of synthetic and natural melanins for a range of inorganic ions showed that zinc was on the lower scale of ionic afﬁnity [130] and the binding afﬁnity of Zn2+ to melanin is less than that of Cu2+[128], melanosomes often contain more zinc than copper [122]. It has been shown that the binding sites for Zn2+ may vary according to pH; at physiological pH mixed complexes of Zn-quinone imine and Zn-catechol dominate [138]. Consistent with this, two classes of independent binding sites participating in the interactions of Zn2+ with dopa-melanin have been demonstrated with association constants of K1 = 5.87 × 105 and K2 = 4.85 × 103 [139]. Non-melanin-binding sites, which include TRP-2 as a zinc-containing enzyme (see Chapter 4) and α-mannosidase as an enzyme potentiated by Zn2+ [140], are likely to be of minor signiﬁcance. There are a number of possible explanations for the high zinc content of melanosomes. It is possible that the metal is implicated in some structural role. Zinc (and to some extent copper and cobalt) ions are capable of inducing structural modiﬁcations in the biosynthesis of dopa-melanins [30]. It is of some interest that zinc, as a diamagnetic ion, enhances the electron paramagnetic resonance signal on binding to melanin due to stabilization of semiquinone free radicals in contrast to the effect of paramagnetic ions such as iron [51]. Two mechanisms of zinc action have been elucidated – the protection of -SH groups against oxidation and the inhibition of the production of ROS by some transition metals, and, on this basis, these authors believe that the essential biochemical function of zinc is to serve as a natural antioxidant [141, 142]. Hence, a high concentration of zinc might be expected in tissues vulnerable to oxidative stress such as spermatozoa, skin, hair, and the eye [142]. Melanosomes, as organelles in which melanogenesis is compartmentalized, makes them a location of oxidative stress and a site of free radical generation (see Chapter 3 and Section 7.6) and might, therefore, beneﬁt from a high concentration of zinc. Another point to be considered is that zinc ions are extremely cytotoxic agents [143] and their sequestration in melanosomes may represent an important mechanism of detoxiﬁcation. In this regard skin and hair melanosomes might participate in zinc excretion [125, 144]. The zinc pool in melanosomes has been shown to be labile: it was possible to remove all radioactive zinc by a 5-day exchange diffusion against 1 mmol/l ZnCl2 from B16 melanoma melanosomes labeled with 65Zn in vivo [135]. Therefore, the protective function of melanosomes is dependent on the permeability barrier provided by the melanosomal membrane that appears to be defective in melanoma cells [65] and it is interesting that melanoma cells are more sensitive to the toxic effects of Zn2+ than normal melanocytes [145]. Similarly, Farmer et al. [146]

12.6 Melanosomes and Metal Ions

reported that the addition of Zn2+ (or Cu2+) to pigment cells in vitro induced more cell death in melanoma cells than in normal melanocytes. Although zinc binds to melanin more strongly than calcium [128], melanosomes may play an important physiological role in relation to the binding of calcium ions. Bush and Simon [134] determined the association constant for Ca2+-binding to Sepia melanin granules to be 3.3 ± 0.2 × 103/mol, which is compatible with buffering or storage of these ions by melanin. Calcium binding by catechols occurs through ionic bonds either through single or adjacent catecholic residues [128, 147], and thus melanosomes containing melanin with signiﬁcant catechol concentration are able to sequester calcium and aid intracellular calcium homeostasis [148]. There seems to be a difference between ocular and epidermal melanosomes; in the pigmented tissues of the eye melanosomes were reported to contain up to 10 times more calcium than the adjacent cytoplasm, whereas this was not the case in the skin [149]. Possibly this difference may be ascribed to the absence of turnover of ocular pigment. As calcium is a key signaling molecule in the orchestration of apoptosis [150] the calcium sequestration by melanosomes may play a crucial role in cytoprotection. Indirect evidence of this action comes from the inﬂuence of inner ear melanin on acoustic injury. It has been shown that the pathogenesis of acoustic damage involves L-type gated calcium channels [151]. It is known that brown-eyed individuals are less prone than blue-eyed individuals to acoustic injury [152] and it has been suggested that melanin may be involved in the control of calcium levels in the endolymph [153, 154]. Similarly, albino mice are sensitive to acoustic injury and, in a series of elegant experiments, Montoliu et al. have shown that, by inducing ectopic expression of tyrosine hydroxylase in the ear melanocytes of transgenic albino mice, the susceptibility to acoustic injury is abrogated [155]. Since the product of tyrosine hydroxylase activity is dopa, the implication is that the generation of this catechol in the melanocytes of the stria vascularis is able to substitute for the absence of melanin in the albino animals by binding calcium and thus have a cytoprotective effect on the hair cells. A further example of this putative effect is the ectopic expression of tyrosine hydroxylase in the pigment epithelium, which rescues the retinal abnormalities and visual function of albinos in the absence of melanin [156]. An interesting corollary of this argument is that if calcium binding by melanin is cytoprotective, then alterations in the state of the melanin may inﬂuence the fate of cells in which the melanosomes are located. One of the properties of melanin is its ability to undergo facile redox change and, in particular, to undergo photo-oxidation. The effect of irradiation is to alter the redox state of melanin and hence modify the catechol/quinone ratio. It follows that the conversion of catecholic to quinone moieties will diminish the sites available for calcium binding. In a model system it has been shown that calcium bound to 4-methylcatechol is released when the catechol is oxidized by tyrosinase (Stratford and Riley, unpublished experiments) and, by analogy, it can be argued that irradiation of melanin would release calcium ions, which on entering the cytosol could initiate apoptosis in cells exposed to sufﬁciently high doses. It follows that this mechanism would

363

364

12 Physiological and Pathological Functions of Melanosomes

be most effective in the most pigmented cells as shown by the greater number of necrotic “sunburn cells” in black compared to white epidermis following equivalent UV exposure and that macrophages loaded with melanin were more susceptible to UV damage [157]. Also, experiments by MacDonald et al. [158] on tricolor guinea pigs showed that the degree of epidermal necrosis was proportional to the degree of pigmentation. If calcium release from melanosomes has a signaling function as proposed, another functional aspect is the possibility of metal ion displacement by agents that bind to melanin. In this connection it has been shown that ototoxicity caused by neomycin is prevented by a calpain inhibitor [159]. One of the prerequisites for calcium ions to act as signaling agents or second messengers is the normal barrier function of cell membranes and, in this regard, it is important to note that during isolation procedures melanosomes usually lose their limiting membranes. For this reason the data derived from experiments with isolated melanosomes or melanins may not necessarily reﬂect the situation in intact cells. In this context it is of interest that Liu et al. [136] reported the existence of channels within Sepia melanin granules that can serve to transport metal ions. Whether this is speciﬁc to cephalopod pigment granules or a general phenomenon of melanosomes remains to be answered.

12.7 Afﬁnity of Melanosomes for Polycyclic and Other Compounds

The afﬁnity of pigmented tissues for phenothiazine derivatives was revealed when tranquillizers were introduced into psychiatric practice and many treated patients developed ocular problems [160]. Later studies documented that the capacity to bind some chemicals is one of the most conspicuous features of melanin (reviewed by Larsson [161]) and melanosomes (reviewed by Borovanský [1]). In addition to binding ions (see Section 12.6), melanin and melanosomes can bind various organic compounds, including polycyclic amines, aromatic hydrocarbons, and heterocyclic drugs. Although not covalently linked to melanin, these compounds are relatively strongly bound and can be slowly released from preformed melanin in melanosomes over long periods, inducing toxic lesions in pigmented tissues, especially those with little or no turnover. In addition, there is the possibility of inducing secondary injury in the adjacent tissues (reviewed by Lindquist [162]). The drawback of long-term retention with pathological consequences in normal tissues becomes an advantage in handling tumors. In particular, when tagged with an appropriate isotope or cytotoxic moiety, the long-term retention of a compound with afﬁnity to melanin might be clinically useful in the diagnosis, monitoring, and therapy of melanoma (reviewed by Larsson [161]). The question of the transport mechanism(s) of radiopharmaceuticals across melanosomal membranes that, unless defective, seem to have only limited permeability in model experiments [163], has not been addressed in any detail apart from

12.7 Afﬁnity of Melanosomes for Polycyclic and Other Compounds

a study by Cavatorta et al. [164] reporting that melanosomal membranes from various sources differ both in their chemical composition and properties. The processes underlying the afﬁnity of melanin for various ligands are complex and have been reviewed by Larsson [161]. An afﬁnity chromatographic study of several drugs towards immobilized melanin conﬁrmed the complexity of the binding behavior [165]. The interaction between anionic sites in melanin (particularly carboxylic groups) and cationic groups (metal ions, protonated amines) is primarily ionic in nature. In melanin interactions with aromatic and polycyclic compounds hydrophobic bonds and van der Waal’s forces may participate. With regard to electron-donating substances, charge-transfer interactions are possible. A charge-transfer complex or electron donor–acceptor complex is an association of two or more molecules in which there is partial transfer of electronic charge. The resulting electrostatic attraction between the electron donor and electron acceptor provides a stabilizing force for the molecular complex. The effect is generated by an excited state best characterized as a weak electron resonance, often producing intense colors characteristic of these complexes – a well-known example being the blue charge-transfer band exhibited by the iodine test for starch. Chargetransfer complexes are of importance in biological systems [166]. Studies of melanin binding have shown that cationic molecules such as chlorpromazine, chloroquine, paraquat, and methylene blue (MTB) form charge-transfer complexes [167–169], and, in humans, charge-transfer complexes of cationic metals and certain drugs with melanin in the skin, the stria vascularis of the ear, and the neuromelanin of the midbrain may be associated with certain pathological conditions. It is a moot point whether charge-transfer complexes contribute to the spectral properties of melanin. The melanin/melanosome afﬁnity of various ligands can be exploited or may have practical impact in several areas as outlined below. 12.7.1 Melanoma Detection and Treatment

Melanoma detection and treatment has already been mentioned above. A number of selective “melanoma seekers” – the thioureylenes – were discovered at the Department of Toxicology in Uppsala. These compounds, such as thiouracil, methimazole, and thiourea (Figure 12.12), and their derivatives, have a favorable speciﬁc feature. They do not bind to preformed melanin, but are incorporated by covalent bonds into nascent melanin during its synthesis, thereby sparing those a)

Figure 12.12

b)

c)

Structural formulae of thiouracil (a), thiourea (b), and methimazole (c).

365

366

12 Physiological and Pathological Functions of Melanosomes a)

b)

Figure 12.13 Structural formulae of MTB (a) and benzamide (b).

tissues in which melanogenesis is low or absent, such as the pigmented tissue of the eye [161, 170]. Labeled derivatives of these compounds are valuable as diagnostic and therapeutic agents [171]. Their 75Se-derivatives, as γ-emitters, are suitable for scintigraphy, whereas 35S-derivatives, as β-emitters, are suitable for radiotherapy. MTB is considered a safe compound that can be used in medicine. MTB is a phenothiazine derivative (Figure 12.13a), and possesses a high afﬁnity for melanin and accumulates preferentially in melanoma cells. Since it is not directly toxic, MTB serves as a useful carrier for radioisotopes and acts as a selectively localized source of radiation. Hence, radiolabeled derivatives of the compound can be used for diagnosis and therapy of disseminated melanoma. [131I]MTB in conjunction with γ-camera imaging has already proved to be a useful tool in clinical studies for the detection of early melanoma dissemination. Astatinated MTB has been shown to be exceptionally efﬁcacious in treating melanoma and preventing its metastatic spread without damaging normal structures when administered systemically to human melanoma-bearing mice. Astatine-211 is an α-emitter with very high linear energy transfer and a phase I clinical trial of [211At]MTB has been approved [172, 173]. Radiolabeled-benzamide derivatives have a long history as melanoma imaging agents [174]. After repeated demonstrations of their selective deposition in melanosomes by analytical imaging using secondary ion mass spectrometry [175, 176], interest in them has persisted up to the present [177]. Similarly, nicotinamide derivatives are currently receiving much attention [178]. 12.7.2 Participation of Melanosomes in Chemoresistance

The melanosomal sequestration of cytotoxic drugs has been noted in the case of cis-diaminedichloroplatinum II (cisplatin) [179] and the acridone derivative [180] with the result that the nuclear localization of these agents is diminished with signiﬁcant chemoresistance of melanoma cells. Moreover, in the case of cisplatin, its melanosomal incorporation induced both melanogenesis and extracellular transport of melanosomes containing cisplatin [179]. Preventing melanosomal sequestration of cytotoxic drugs by inhibiting the functions of melanosomes may

12.7 Afﬁnity of Melanosomes for Polycyclic and Other Compounds

have great potential as an approach to improving the chemosensitivity of melanoma cells. Melanoma drug sensitivity has been reported to be inﬂuenced by melanosome dynamics: melanosomes of stage II and III exhibited the highest potency in decreasing sensitivity to cisplatin by drug trapping. Interventions that alter the numbers and stages of melanosomes could, therefore, provide a means of increasing cellular sensitivity to chemotherapeutic agents [181]. A similar study by Xie et al. [182], also using cisplatin, gave the ﬁrst direct evidence that melanosomal regulatory genes inﬂuence drug sensitivity. Absence of melanosomal structural protein gp100/Pmel17, independent mutations in three separate genes regulating biogenesis of melanosomes (DTNBP1, PLDN, and VPS33A), and a mutation in the gene coding for tyrosinase induced increased cisplatin sensitivity. Sensitivity to other chemotherapeutics, including vinblastine and etoposide, also increased with mutation of PLDN. The presence of mature melanosomes was considered to contribute to melanoma resistance to chemotherapy – a result at variance with the study [181] in which stage IV melanosomes were reported to increase drug sensitivity. 12.7.3 Long-Term Deposition of Compounds in Melanosomes

Melanosomes, especially those incorporated into keratinized structures such as hair, can retain various compounds, ions, and drugs for long periods. Using tritiated cocaine, nicotine, and ﬂunitrazepam, their deposition in hair melanosomes after systemic administration has been demonstrated [183]. This can be of practical use in forensic pathology, in detecting drug abuse, and in pollution monitoring. Explosives [184], methadone [185], and codeine [186] were always found in the highest concentration in black hair. Experiments have been performed to determine the in vitro binding of cocaine (Figure 12.14), benzoylecgonine (BE), amphetamine, and N-acetylamphetamine (N-AcAp) to synthetic melanin subtypes. The melanins investigated included two eumelanin subtypes, based on DHI and DHICA, a reddish-brown pheomelanin, derived from 5-S-cysteinyldopa, and two mixed eu/pheomelanin copolymers. The results showed that the basic compounds, cocaine and amphetamine, bind to eumelanins and mixed eu/pheomelanins to varying degrees, but not to pure pheomelanin. BE and N-AcAp, net neutral molecules, fail to bind to any type of melanin. Further studies, using tandem mass spectrometry, on amphetamine

Figure 12.14

Structural formula of cocaine.

367

368

12 Physiological and Pathological Functions of Melanosomes

binding to eumelanin showed noncovalent binding to quinonoid dimers. Similar functional groups on the eumelanin polymer may represent an important drug-binding site. Overall, these ﬁndings show that basic drugs have a greater afﬁnity for melanin than their net neutral analogs and reveal that melanin types differ when it comes to drug binding, thus helping to explain why hair color biases exist [187].

12.8 Exploitation of Melanosomal Proteins and Melanin as Speciﬁc Targets in Melanoma Therapy

All the above-mentioned properties and functions of melanosomes and strategies of their exploitation in medicine have been derived from the presence of melanin in melanosomes or from the process of melanogenesis. However, even if melanosomes are poorly melanized and melanogenesis is reduced or absent, melanosomes possess features that can be of practical use. The introduction of monoclonal antibodies has led to a rapid advance in information concerning not only the antigenic structures of melanoma cell membranes, but also those of melanosomes [1]. The evidence for the presence of antigenic determinants common to the inner melanosomal compartment and to the cell surface of pigment cells [188] has opened a gateway to their exploitation for immunodiagnostic tests and immunotherapy of malignant melanoma. The range of melanosomal proteins is detailed in Chapters 9 and 11. Many of them, including epitopes derived from tyrosinase, TRP-1/gp75, Pmel17/silver/ gp100, and MART-1/Melan-A, function in vivo as targets for humoral and cellular autoimmune responses directed against normal and/or transformed pigment cells. Hence, melanosomes can be considered as a speciﬁc source of immune targets for melanoma [189, 190]. In their review, Marks et al. [191] underlined that mechanisms that regulate the biogenesis of melanosomes, and the pathways by which constituent proteins are targeted to them, are related to those involved in the biogenesis of the major histocompatibility complex (MHC) class II antigen processing compartment. Many of the tissue-speciﬁc proteins that localize to melanosomes and participate in melanin formation double as tumor-associated antigens that are targets for T cells in patients with melanoma. Melanosomes at different stages of development are enriched in distinct protein pools. Mechanisms regulating the loading and intracellular trafﬁcking of MHC class II molecules for successful antigen presentation have been recently summarized by Rocha and Neefjes [192]. An important mechanism in inducing an immunological response is protein haptenation (i.e., the chemical modiﬁcation of self-protein(s)), thus forming macromolecular immunogens [193]. Reactive o-quinones generated by tyrosinasecatalyzed oxidation in additive reaction with nucleophilic functions such as amino or thiol group can induce haptenation [194]. The most proximal haptenation

12.8 Exploitation of Melanosomal Proteins and Melanin as Speciﬁc Targets in Melanoma Therapy

targets are intramelanosomal proteins and it is noteworthy that the melanosome contains the majority of the antigenic proteins of melanocytes [190]. As the protein nearest to the source of o-quinones, tyrosinase is especially vulnerable to haptenation. In the case of 4-hydroxyanisole it has been demonstrated by radiolabeling techniques that the oxidation product binds to tyrosinase [195]. Other potential intramelanosomal haptenation targets include TRP-1, MART-1, Pmel17, and TRP2. In the case of the HLA-A2 isotype, there are some data relating to MHC class I peptides generated from melanosomal proteins. In fact, haptenation has been suggested as a means of enhancing antigenicity by testing reactivity of MART-1reactive T cell clones after modiﬁcation with quinone methide. The antigenic melanosomal protein Melan-A/MART-1 exhibits an immunodominant epitope for HLA-A*0201, EAAGIGILTV, which has been identiﬁed by cytotoxic T lymphocyte screening [196]. Clones reactive to the native peptide maintained reactivity after nucleophilic addition affecting the thiol side group of cysteine [197]. Further studies are needed to identify T cell reactivity elicited speciﬁcally by haptenated peptides presented in the context of HLA-A2 and other human leukocyte antigen (HLA) isotypes. The susceptibility of developing an immune response to a neoantigen generated by a haptogenic agent is inﬂuenced by the efﬁciency with which antigenic peptides are presented to the immune system [198]. Consequently, the HLA isotype expressed by individuals will contribute to the likelihood of developing an immune response. Consistent with this interpretation, Morgan et al. [199]. reported a clinical pilot study in which 4-hydroxyanisole was administered intra-arterially to a group of melanoma patients with widespread disease unresponsive to other therapies. Despite the high doses administered, the acute responses were disappointing, but longer-term follow-up showed that 45% of cases showed some degree of regression by tumors in the infused zone, although there was no evidence of a generalized tumor response or of any obvious cutaneous or ocular depigmentation. In one case there was complete regression of secondary tumors. This patient, with multiple recurrences at the site of a skin graft on the right leg, received two courses of 4-hydroxyanisole (100 and 84 g by intra-femoral infusion) with a 4-week interval and, when seen 4 weeks after the second infusion, was tumor-free [200]. This delayed reaction would be explained by the initiation of an immune response speciﬁc to tyrosinase-expressing cells exposed to 4-hydroxyanisole [201, 202]. A similar process is probably reﬂected in Lipizzaner and Camargue horses as well as Sinclair swine, where gradual depigmentation is a consequence of melanomas “spontaneously” regressing due to an immune response. Preparation of monoclonal antibodies to melanin has emerged as an attractive strategy for targeting radioimmunotherapy of melanoma. Monoclonal antibodies 6D2 and 11B11 tagged with 188Rc were shown to have higher therapeutic efﬁcacy compared to dacarbazine chemotherapy in human melanoma xenografts in nude mice [203], and it is likely that melanin-targeted radionuclide therapy will enter clinical practice within the next decade [204].

369

370

12 Physiological and Pathological Functions of Melanosomes

12.9 Conclusions

Many of the properties ascribed to melanosomes are those deriving from the characteristics of melanins, such as their optical absorbance, their redox properties, and their propensity for binding cationic materials, including drugs and metals [205]. However, their existence as specialized organelles is based on the requirement for cells to contain the hazardous process of melanogenesis. From an evolutionary standpoint it is probable that the initial advantage arising from the expression of an enzyme able to generate reactive o-quinones would consist of utilization of their toxic properties as a defensive mechanism, such a role being served by a secretory pathway. However, the remarkable light-absorbing properties of indolic products arising from the oxidation of phenolamine substrates, and the physiological advantages arising from this, may have been the stimulus to retain the pigment and thus internalize the process of melanin formation. This necessitates the compartmentalization of the process to prevent widespread cytotoxic damage by the reactive intermediates of oxidation, and also the provision of a framework on which the pigment can be deposited and retained. The outcome of this evolutionary pressure is the melanosome – an organelle whose complex biogenesis and distribution is set out in detail in Chapters 9 and 10. The crucial importance of limiting the cellular location both of the reactants and the pigment is illustrated by the damage associated with uncontained melanin as in the case of neuromelanin, which arises by a different pathway (as described in Chapter 8). As well as limiting the pathological consequences of cytosolic diffusion of melanins and melanogens, melanosomes may serve important homeostatic functions by regulating ion exchange with the cytosol, and the integrity of melanosomal membranes and associated proteins is of great physiological and pathological signiﬁcance. Finally, there are differences in the roles of melanosomes that are retained (as in the eye and ear, see Chapters 2 and 7) and those that are distributed to cells other than melanocytes, and which exhibit a turnover. Cells that retain melanosomes are at greater risk of long-term toxicity by agents with pronounced melanin afﬁnity, while melanosomes that are distributed to exfoliating cells (as in the epidermis) may act as an excretory pathway for toxins. Clearly the melanosome is a remarkable and unique organelle, and we may yet ﬁnd that it serves further important and hitherto undiscovered biological functions.

Acknowledgments

Supported by VZ MSTM CR 21620808.

References

References 1 Borovanský, J. (1993) Properties of

2

3

4

5

6

7

8

9

10

melanosomes and their exploitation in the diagnosis and treatment of melanoma. Melanoma Res., 3, 181–186. Ito, S. and Wakamatsu, K. (2003) Quantitative analysis of eumelanin and pheomelanin in humans, mice, and other animals: a comparative review. Pigment Cell Res., 16, 523–531. Stamatas, G.N., Zmudzka, B.Z., Kollias, N., and Beer, J.Z. (2004) Non-invasive measurements of skin pigmentation in situ. Pigment Cell Res., 6, 18–26. Yudovsky, D. and Pilon, L. (2010) Rapid and accurate estimation of blood saturation, melanin content, and epidermal thickness from spectral diffuse reﬂectance. Appl. Opt., 49, 1707–1719. Nielsen, K.P., Zhao, L., Ryzhikov, G.A., Biruyulina, M.S., Sommersten, E.R., Stamnes, J.J., Stamnes, K., and Moan, J. (2008) Retrieval of the physiological state of human skin from UV-VIS reﬂectance spectra – a feasibility study. J. Photochem. Photobiol B, 93, 23–31. Heymann, J.A., Shi, D., Kim, S., Bliss, D., Milne, J.L., and Subramaniam, S. (2009) 3D imaging of mammalian cells with ion-abrasion scanning electron microscopy. J. Struct. Biol., 166, 1–7. Borovanský, J., Vedralová, E., and Hach, P. (1991) An estimate of melanosome concentration in pigment tissues. Pigment Cell Res., 4, 222–224. Duchon, J., Borovanský, J., and Hach, P. (1973) Chemical composition of ten kinds of various melanosomes, in Mechanisms in Pigmentation (eds V.J. McGovern and P. Russell), Karger, Basel, pp. 165–170. Borovanský, J. and Duchon, J. (1974) Chemical composition of hair melanosomes. Dermatologica, 149, 116–120. Borovanský, J., Horák, V., Elleder, M., Fortýn, K., Smit, N.P., and Kolb, A.M. (2003) Biochemical characterization of a new melanoma model – the minipig MeLiM strain. Melanoma Res., 13, 543–548.

11 Seiji, M. and Shimao, K. (1961) [Density

12

13

14

15

16

17

18

19

gradient method]. J. Jap. Biochem. Soc., 33, 435–441. Jimbow, K., Miyake, Y., Homma, K., Yasuda, K., Izumi, Y., Tsutsumi, A., and Ito, S. (1984) Characterization of melanogenesis and morphogenesis of melanosomes by physicochemical properties of melanin and melanosomes in malignant melanoma. Cancer Res., 44, 1126–1134. Vedralová, E., Borovanský, J., and Hach, P. (1995) Ganglioside proﬁles of experimental melanomas and of their melanosomal fractions. Melanoma Res., 5, 87–92. Vedralová, E. and Duchon, J. (1983) Comparison of lipids between tumor and normal hamster melanosomes. Neoplasma, 30, 317–321. Diment, S., Eidelman, M., Rodriguez, G.H., and Orlow, S.J. (1995) Lysosomal hydrolases are present in melanosomes and are elevated in melaninzing cells. J. Biol. Chem., 270, 4213–4215. Borovanský, J. and Elleder, M. (2003) Melanosome degradation: fact or ﬁction? Pigment Cell Res., 16, 280–286. Yates, T.J.R., III, Gilchrest, A., Howell, K.E., and Bergeron, J.J.M. (2005) Proteomics of organelles and large cellular structures. Nat. Rev. Mol. Cell. Biol., 6, 702–714. Basrur, V., Yang, F.G., Kushimoto, T., Higashimoto, Y., Yasumoto, K., Valencia, J., Muller, J., Vieira, W.D., Watabe, H., Shabanowitz, J., Hearing, V.J., Hunt, D.F., and Appella, E. (2003) Proteomic analysis of early melanosomes: identiﬁcation of novel melanosomal proteins. J. Proteome Res., 2, 69–79. Chi, A., Valencia, J.C., Hu, Z.Z., Watabe, H., Yamaguchi, H., Mangini, N.J., Husang, H., Canﬁeld, V.A., Cheng, K.C., Yang, F., Abe, R., Yamagishi, S., Shabanowitz, J., Hearing, V.J., Wu, C., Appella, E., and Huint, D.F. (2006) Proteomic and bioinformatic characterization of the biogenesis and

371

372

12 Physiological and Pathological Functions of Melanosomes

20

21

22

23

24

25

26

27

28

29

30

function of melanosomes. J. Proteome Res., 5, 3135–3144. Hu, Z.Z., Valencia, J.C., Huang, H., Chi, A., Shabanowitz, J., Hearing, V.J., Appella, E., and Wu, C. (2007) Comparative bioinformatics analyses band proﬁling of lysosome-related organelle proteomes. Int. J. Mass Spectrom., 259, 147–116. Zajac, G.W., Gallas, J.M., Cheng, J., Eisner, M., Moss, S.C., and AlvaradoSwaisgood, A.E. (1994) The fundamental unit of synthetic melanin: a veriﬁcation by tunneling microscopy of X-ray scattering results. Biochem. Biophys. Acta, 1199, 271–278. d’Ischia, M., Napolitano, A., Pezzella, A., Land, E.J., Ramsden, C.A., and Riley, P.A. (2005) 5,6-Dihydroxyindoles and indole-5,6-diones. Adv. Hetrocycl. Chem., 289, 1–63. Meredith, P. and Sarna, T. (2006) The physical and chemical properties of eumelanin. Pigment Cell Res., 19, 572–594. Borovanský, J. and Duchon, J. (1975) Comparative study of the amino acid composition of some tumor and normal melanosomes. Neoplasma, 22, 195–199. Chedekel, M., Ahene, A.B., and Zeise, L. (1992) Melanin standard method: empirical formula 2. Pigment Cell Res., 5, 240–246. Wagh, S., Ramaiah, A., Subramanian, R., and Govindajaran, R. (2000) Melanosomal proteins promote melanin polymerization. Pigment Cell Res., 13, 442–448. Vohra, F. and Kratzer, F.H. (1959) Incorporation of d,l-lysine-2-14C into melanin pigment of turkey poult feathers. Proc. Soc. Exp. Biol. Med., 100, 837–838. Yasunobu, K.T., Peterson, E.W., and Mason, H.S. (1959) The oxidation of tyrosine-containing peptides by tyrosinase. J. Biol. Chem., 234, 3291–3295. Ito, S., Kato, T., Shinpo, K., and Fujita, K. (1984) Oxidation of tyrosine residues in proteins by tyrosinase. Biochem. J., 222, 402–411. Palumbo, A., Ischia, M., Misuraca, G., Prota, G., and Schultz, T.M. (1988) Structural modiﬁcations in biosynthetic

31

32

33

34

35

36

37

38

39

40

41

42

melanins induced by metal ions. Biochim. Biophys. Acta, 964, 193–199. Kempf, V.R., Wakamatsu, K., Ito, S., and Simon, J.D. (2010) Imaging, chemical and spectroscopic studies of methylation-induced decomposition of melanosomes. Photochem. Photobiol., 86, 765–771. Sharma, A. (2010) Effect of ambient humidity on UV/visible photodegradation of melanin thin ﬁlms. Photochem. Photobiol., 86, 852–855. Hach, P., Borovanský, J., and Vedralová, E. (1993) Melanosome – a sophisticated organelle. Sborník. Lék., 94, 113–123. Sakurai, T., Ochia, I., Takeuchi, H., and Jimbow (1975) Ultrastructural change of melanosomes associated with agouti pattern formation in mouse hair. Dev. Biol., 47, 466–471. T., K., Takeuchi and T. (1979) Ultrastructural comparison of pheo- and eumelanogenesis in animals. Pigment Cell, 4, 308–317. Ito, S. and Fujita, K. (1985) Microanalysis of eumelanin and pheomelanin in hair and melanomas by chemical degradation and liquid chromatography. Anal. Biochem., 144, 527–536. Stanka, P. (1974) Ultrastructural study of pigment cells of human red hair. Cell Tissue Res., 150, 167–178. Hach, P. and Borovanský, J. (1972) Ultrastructure of melanosomes of different origin. Folia Morphol., 20, 82–84. Borovanský, J., Hach, P., and Duchon, J. (1977) Melanosome: an unusually resistant subcellular particle. Cell Biol. Int. Rep., 1, 549–552. Moyer, F.H. (1966) Genetic variations in the ﬁne structure and ontogeny of mouse melanin granules. Am. Zool., 6, 43–66. Borovanský, J., Mommaas, A.M., Smit, N.P.M., Eygendaal, D., Winder, A.J., Vermeer, B.J., and Pavel, S. (1997) Melanogenesis in transfected ﬁbroblasts induces lysosomal activation. Arch. Dermatol. Res., 289, 145–150. Sealy, R.C. (1984) Free radicals in melanin formation, structure and reactions, in Free Radicals in Molecular

References

43

44

45

46

47

48

49

50

51

52

53

Biology, Aging, and Disease (eds D. Armstrong, R.S. Sohal, R.G. Cutler, and T.F. Slater), Raven Press, New York, pp. 67–76. McGinness, J.E., Kono, R., and Moorhead, W.D. (1970) The melanosome: cytoprotective or cytotoxic? Pigment Cell, 4, 270–276. Borovanský, J. (1996) Free radical activity of melanins and related substances: biochemical and pathobiochemical aspects. Sborník. Lék., 97, 49–70. Chedekel, M. (1982) Photochemistry and photobiology of epidermal melanins. Photochem. Photobiol., 35, 881–885. Ezzahir, A. (1989) The inﬂuence of melanins on peroxidation of lipids. J. Photochem. Photobiol. B, 3, 341–349. Halliwell, B. (1995) Antioxidant characterization: methodology and mechanism. Biochem. Pharmacol., 49, 1341–1348. Schweitzer, A.D., Revskaya, E., Chu, P., Pazo, V., Friedman, M., Nosanchik, J..D., Cahill, S., Frases, S., Casadevall, A., and Dadachova, E. (2010) Melanincovered nanoparticles for protection of bone marrow during radiation therapy of cancer. Int. J. Radiat. Oncol. Biol. Phys., 78, 1494–1502. Simon, J.D. and Peles, D.N. (2010) The red and the black. Acc. Chem. Res., 43, 1452–1460. Riley, P.A. (1977) The mechanism of melanogenesis. Symp. Zool. Soc. Lond., 39, 77–95. Sarna, T. and Swartz, H.A. (2006) The physical properties of melanin, in The Pigmentary System: Physiology and Pathophysiology, 2nd edn (eds J.J. Nordlund, R.E. Boissy, V.J. Hearing, R.A. King, W.S. Oetting, and J.P. Ortonne), Blackwell, Malden, MA, pp. 311–341. Felix, C.C., Hyde, J.S., Sarna, T., and Sealy, R.C. (1978) Melanin photoreactions in aerated media. Electron spin resonance evidence for production of superoxide and hydrogen peroxide. Biochem. Biophys. Res. Commun., 84, 335–341. Sarna, T., Duleba, A., Korytowski, W., and Swartz, H. (1980) Interaction of

54

55

56

57

58

59

60

61

62

63

melanin with oxygen. Arch. Biochem. Biophys., 200, 140–148. Ranadive, N.S. and Menon, I.A. (1986) Role of reactive oxygen species and free radicals from melanins in photoinduced cutaneous inﬂammations. Pathol. Immunopathol. Res., 5, 118–139. Korytowski, W., Pilas, B., Sarna, T., and Kalyanamaran, B. (1987) Photoinduced generation of hydrogen peroxide and hydroxyl radicals in melanomas. Photochem. Photobiol., 45, 185–190. Bittinger, F., Gonzáles-García, J.L., Klein, C.L., Brockhausen, C., Offner, F., and Kirckpatrick, C.J. (1998) Production of superoxide dismutase by human melanoma cells. Melanoma Res., 8, 381–387. Elleder, M. and Borovanský, J. (2001) Autoﬂuorescence of melanins induced by ultraviolet radiation and near ultraviolet light. A histochemical and biochemical study. Histochem. J., 33, 273–281. Menon, I.A., Persad, S., Ranadive, N.S., and Haberman, H.F. (1983) Effects of ultraviolet-visible irradiation in the presence of melanin isolated from human black or red hair upon Ehrlich ascites carcinoma cells. Cancer Res., 43, 3165–3169. Bouchilloux, S. and Kodja, A. (1960) Combinaison des thiols avec les quinones se formant au cours de la mélanogénèse. Bull. Soc. Chim. Biol., 42, 1045. Miranda, M., Botti, D., and DiCola, M. (1984) Possible genotoxicity of melanin synthesis intermediates. Tyrosinase reaction products interact with DNA in vitro. Mol. Gen. Genet., 193, 395–399. Miranda, M., Amicarelli, F., Poma, A., Ragnelli, A., Scirri, C., Aimola, P.P., Masciocco, L., Bonﬁgli, A., and Zarivi, O. (1994) Cytogenotoxic species leakage with human melanoma melanosomes. Molecular–morphological correlations. Biochem. Mol. Biol. Int., 32, 913–922. Riley, P.A. (1988) Radicals in melanin biochemistry. Ann. NY Acad. Sci., 551, 111–120. Riley, P.A. (1970) Mechanism of pigment cell toxicity produced by hydroxyanisole. J. Pathol., 101, 163–169.

373

374

12 Physiological and Pathological Functions of Melanosomes 64 Borovanský, J., Mirejovsky, P., and Riley,

65

66

67

68

69

70

71

72

73

74

P.A. (1991) Possible relationship between abnormal melanosome structure and cytotoxic phenomena in malignant melanoma. Neoplasma, 38, 393–400. Hunter, J.A.A., Paterson, W.D., and Fairley, D.J. (1978) Human malignant melanoma. Melanosomal polymorphism and the ultrastructural dopa reaction. Br. J. Dermatol., 98, 381–390. Curran, R.C. and McCann, B.G. (1976) The ultrastructure of benign pigmented naevi and melanocarcinomas in man. J. Pathol., 119, 135–146. Klingmüller, G. and Schmoeckel, C. (1971) Frei im Cytoplasma liegende Melanin-synthesierende Membranordnungen beim malignen Melanom. Arch. Derm. Forsch., 241, 115–121. Hochstein, P. and Cohen, G. (1963) The cytotoxicity of melanin precursors. Ann. NY Acad. Sci., 100, 876–886. Offner, F.A., Wirtz, H.C., Schiefer, J., Bigalke, I., Klosterhalfen, B., Bittinger, F., Mittemayer, C., and Kirkpatrick, C.J. (1992) Interaction of human malignant melanoma (ST-ML-12) tumor spheroids with endothelial cell monolayers. Damage to endothelium by oxygenderived free radicals. Am. J. Pathol., 141, 601–610. Lapina, V., Dontsov, A.E., and Ostrovskij, M.A. (1984) [Superoxide generation via melanin interaction with oxygen]. Biochimija, 49, 1712–1718. Carstam, R., Edner, C., Hansson, C., Lindbladh, C., Rorsman, H., and Rosengren, E. (1986) Metabolism of 5-S-glutathione-dopa. Acta Derm. Venereol. Suppl., 66, 1–12. Pawelek, J.M. and Lerner, A.B. (1978) 5,6-Dihydroxyindole is a melanin precursor showing potent cytotoxicity. Nature, 276, 627–628. Pavel, S. and Smit, N.P.M. (1996) Metabolic interference of melanogenesis in pigment cells. Sborník. Lék., 97, 29–39. Duchon, J. (1987) Urinary melanogens as a mirror of melanogenesis in vivo, in Cutaneous Melanoma (eds U. Veronesi, N. Cascinelli, and N. Santinami),

75

76

77

78

79

80

81

82

83

84

Academic Press, London, pp. 225–232. Agrup, G., Agrup, P., Andersson, T., Hafström, L., Hansson, C., Jacobsson, S., Jönsson, P.E., Rorsman, H., Rosengren, A.M., and Rosengren, E. (1979) 5 Years’ experience of 5-Scysteinyldopa in melanoma diagnosis. Acta Derm. Venereol., 59, 381–388. Takahashi, H. and Fitzpatrick, T.B. (1966) Large amounts of dihydroxyphenylalanine in the hydrolysate of melanosomes from Harding-Passey mouse melanoma. Nature, 209, 888–890. Rodgers, K.J. and Dean, R.T. (2000) Metabolism of protein-bound DOPA in mammals. Int. J. Biochem. Cell Biol., 32, 945–955. Rodgers, K.J., Hume, P.M., Dunlop, R.A., and Dean, R.T. (2004) Biosynthesis and turnover of DOPA-containing proteins by human cells, Free Rad. Biol. Med., 37, 1756–1764. Ito, S., Kato, K., Shinpo, K., and Fujita, K. (1984) Oxidation of tyrosine residues in proteins by tyrosinase. Formation of protein-bonded 3,4-dihydroxyphenylalanine and 5-S-cysteinyl-3,4-dihydroxyphenylalanine. Biochem. J., 222, 407–411. Nelson, M., Foxwell, A..R., Tyrer, P., and Dean, R.T. (2007) Protein-bound 3,4-dihydroxyphenylalanine (DOPA), a redox-active product of protein oxidation, as a trigger for antioxidant defences. Int. J. Biochem. Cell Biol., 39, 879–889. Nelson, M., Foxwell, A.R., Tyrer, P., and Dean, R.T. (2010) Radical sequestration by protein-bound 3,4-dihydroxyphenylalanine. Int. J. Biochem. Cell Biol., 42, 755–761. Borovanský, J., Pavel, S., Duchon, J., and Vulterin, K. (1979) Incorporation of l-3,4-dihydroxy-[2-C14]phenylalanine into hamster melanoma melanosomes. FEBS Lett., 104, 291–293. Morin, B., Davies, M.J., and Dean, R.T. (1998) The protein oxidation product 3,4-dihydroxyphenylalanine (DOPA) mediates oxidative DNA damage. Biochem. J., 330, 1059–1067. Smit, N.P.M., Pavel, S., and Riley, P.A. (2000) Mechanisms of control of the

References

85

86

87

88

89

90

91

92

93

cytotoxicity of orthoquinone intermediates of melanogenesis, in Role of Catechol Quinone Species in Cellular Toxicity (eds C.R. Creveling), F.P. Graham, Johnson City, TN, pp. 191–245. Smit, N.P.M., Van Nieuwpoort, F.A., Marrot, L., Out, C., Poorthuis, B., Van Pelt, H., and Pavel, S. (2008) Increased melanogenesis is a risk factor for oxidative DNA damage – a study on cultured melanocytes and atypical nevus cells. J. Invest. Dermatol., 84, 550–555. Riley, P.A. (1991) Melanogenesis: a realistic target for antimelanoma therapy? Eur. J. Cancer, 27, 1172–1177. Thompson, A., Land, E.J., Chedekel, M.R., Subbarao, K.V., and Truscott, T.G. (1985) A pulse radiolysis investigation of the oxidation of the melanin precursors 3,4-dihydroxyphenylalanine (dopa) and the cysteinyldopas. Biochem. Biophys. Acta, 843, 49–57. Riley, P.A. (1969) Hydroxyanisole depigmentation; in vitro studies. J. Pathol., 97, 185–191. Manini, P., Napolitano, A., Westerhof, W., Riley, P.A., and d’Ischia, M. (2009) A reactive ortho-quinone generated by tyrosinase-catalyzed oxidation of the skin-depigmenting agent monobenzone: self-coupling and thiol-conjugation reactions and possible implications for melanocyte toxicity. Chem. Res. Toxicol., 22, 1398–1405. Land, E.J., Cooksey, C.J., and Riley, P.A. (1990) Reaction kinetics of 4-methoxy ortho benzoquinone in relation to its mechanism of cytotoxicity: a pulse radiolysis study. Biochem. Pharmacol., 39, 1133–1135. Riley, P.A., Cooksey, C.J., Johnson, C.I., Land, E.J., Latter, A.M., and Ramsden, C.A. (1997) Melanogenesis-targeted anti-melanoma pro-drug development: effect of side-chain variations on the cytotoxicity of tyrosinase-generated ortho-quinones in a model screening system. Eur. J. Cancer, 33, 135–143. Oliver, E.A., Schwartz, L., and Warren, L.H. (1940) Occupational leukoderma. Arch. Dermal., 42, 993–998. Riley, P.A. (1974) Disorders of melanin metabolism, in The Physiology and Pathophysiology of the Skin, vol. 3 (ed. A.

94

95

96

97

98

99

100

Jarrett), Academic Press, London, pp. 1157–1189. Gili, A., Thomas, P.D., Ota, M., and Jimbow, K. (2000) Comparison of in vitro cytotoxicity of N-acetyl and N-propionyl derivatives of phenolic thioether amines in melanoma and neuroblastoma cells and the relationship to tyrosinase and tyrosine hydroxylase enzyme activity. Melanoma Res., 10, 9–15. Sato, A., Tamura, Y., Sato, N., Yamashita, T., Takada, T., Sato, M., Osai, Y., Okura, M., Ono, I., Ita, A., Honda, H., Wakamatsu, K., Ito, S., and Jimbbow, K. (2010) Melanoma-targeted chemo-thermo-immuno (CTI)-therapy using N-propionyl-4-Scysteaminylphenol-magnetite nanoparticles elicits CTL response via heat shock protein–peptide complex release. Cancer Sci., 10, 1936–1946. Jordan, A.M., Khan, T.H., Osborn, H.M., Photiou, A., and Riley, P.A. (1999) Melanocyte-directed enzyme prodrug therapy (MDEPT): development of a targeted treatment for malignant melanoma. Bioorg. Med. Chem., 7, 1775–1780. Jordan, A.M., Khan, T.H., Malkin, H., Osborn, H.M.I., Photiou, A., and Riley, P.A. (2001) Melanocyte-directed enzyme prodrug therapy (MDEPT): development of a second generation of prodrugs for the targeted treatment of malignant melanoma. Bioorg. Med. Chem., 9, 1549–1558. Jordan, A.M., Khan, T.H., Malkin, H., and Osborn, H.M.I. (2002) Synthesis and analysis of urea and carbamate prodrugs as candidates for MDEPT. Bioorg. Med. Chem., 10, 2625–2633. Knaggs, S., Malkin, H., Osborn, H.M.I., Williams, N.A.O., and Yaqoob, P. (2005) New prodrugs derived from 6-aminodopamine and 4-aminophenol as candidates for melanocyte-directed enzyme prodrug therapy (MDEPT). Org. Biomol. Chem., 3, 4002–4010. Jawaid, S., Khan, T.H., Osborn, H.M.I., and Williams, N.A. (2009) Tyrosinase activated melanoma prodrugs. Anticancer Agents Med. Chem., 9, 717–727.

375

376

12 Physiological and Pathological Functions of Melanosomes 101 Borovanský, J., Edge, R., Land, E.J.,

102

103

104

105

106

107

108

109

110

Navaratnam, S., Pavel, S., Ramsden, C.A., Riley, P.A., and Smit, N.P.M. (2006) Mechanistic studies of melanogenesis: the inﬂuence of N-substitution on dopamine quinone cyclization. Pigment Cell Res., 19, 170–178. Gasowska –Baier, B., and Wojtasek, H. (2008) Indirect oxidation of the antitumour agent procarbazine by tyrosinase – possible application in designing anti-melanoma prodrugs. Bioorg. Med. Chem. Lett., 18, 3296–3300. Osborn, H.M.I. and Williams, N.A.O. (2004) Development of tyrosinase labile protecting groups for amines. Org. Lett., 6, 3111–3113. Perry, M.J., Mendes, E., Simplicio, A.L., Coelho, A., Soares, R.V., Iley, J., Moreira, R., and Francisco, A.P. (2009) Dopamine- and tyramine-based derivatives of triazenes: activation by tyrosinase and implications for prodrug design. Eur. J. Med. Chem., 44, 3228–3234. Chedekel, M.R. (1995) Photophysics and photochemistry of melanin, in Melanin: Its Role in Human Photoprotection (eds L. Zeise, M.R. Chedekel, and T.B. Fitzpatrick), Valdenmar, Overland Park, KS, pp. 11–22. Riesz, J., Gilmore, J., and Meredith, P. (2006) Quantitative scattering from melanin solutions. Biophys. J., 90, 1–8. Meredith, P. and Riesz, J. (2004) Radiative relaxation quantum yields for synthetic eumelanin. Photochem. Photobiol., 79, 211–216. Goodman, G. and Bercovich, D. (2008) Melanin directly converts light for vertebrate metabolic use: heuristic thoughts on birds, Icarus and dark human skin. Med. Hypotheses, 71, 190–202. Glickman, R.D. (2002) Phototoxicity to the retina: mechanisms of damage. Int. J. Toxicol., 21, 473–490. Faraggi, E., Gerstman, B.S., and Sun, J. (2005) Biophysical effects of pulsed lasers in the retina and other tissues containing strongly absorbing particles: shockwave and explosive bubble generation. J. Biomed. Opt., 10, 240001.

111 Neumann, J. and Brinkmann, R. (2005)

112

113

114

115

116

117

118

119

120

Boiling nucleation on melanosomes and microbeads transiently heated by nanosecond and microsecond laser pulses. J. Biomed. Opt., 10, 06429. Roegener, J., Brinkmann, R., and Lin, C.P. (2004) Pump-probe detection of laser-induced microbubble formation in retinal pigment epithelium cells. J. Biomed. Opt., 9, 367–371. Brinkmann, R., Roider, J., and Birngruber, R. (2006) Selective retina therapy (SRT): a review on methods, techniques, preclinical and ﬁrst clinical results. Bull. Soc. Belge Ophtalmol., 302, 51–69. Framme, C., Walter, A., Prahs, P., Theisen-Kunde, D., and Brinkmann, R. (2008) Comparison of threshold irradiances and online dosimetry for selective retina treatment (SRT) in patients treated with 200 nanoseconds and 1.7 microsecond laser pulses. Lasers Surg. Med., 40, 616–624. Jacques, S.L. and McAuliffe, D.J. (1991) The melanosome: threshold temperature for explosive vaporization and internal absorption coefﬁcient during pulsed laser irradiation. Photochem. Photobiol., 53, 769–775. Wang, D.S. (2009) Intracellular hyperthermia: nanobubbles and their biomedical applications. Int. J. Hypertherm., 25, 533–541. Skrabalak, S.E., Au, L., Lu, X., Li, X., and Xia, Y. (2007) Gold nanocages for cancer detection and treatment. Nanomedicine, 2, 657–668. Lapotko, D. (2009) Optical excitation and detection of vapor bubbles around plasmonic nanoparticles. Opt. Express, 17, 2538–2556. Schweitzer, A.D., Howell, R.C., Jiang, Z., Bryan, R.A., Gerfen, G., Chen, C.C., Mah, D., Cahill, S., Casadevall, A., and Dadachova, A. (2009) Physico-chemical evaluation of rationally designed melanins as novel nature-inspired radioprotectors. PLoS ONE, 4, e7229. Kalyanaraman, B., Felix, C.C., and Sealy, R.C. (1984) Photoionization and photolysis of melanins: an electron spin resonance-spin study. J. Am. Chem. Soc., 106, 7327–7330.

References 121 Lyttkens, L., Larsson, B., Goller, H.,

122

123

124

125

126

127

128

129

130

131

132

133

Englesson, S., and Stahle, J. (1979) Melanin capacity to accumulate drugs in the internal ear – study on lidocaine, bupivacaine and chlorpromazine. Acta Oto-Laryngol., 88, 61–73. Horčičko, J., Borovanský, J., Duchon, J., and Procházková, B. (1973) Distribution of zinc and copper in pigment tissues. Hoppe-Seyler’s Z. Physiol. Chem., 354, 203–204. Larson, B. and Tjälve, H. (1978) Studies on the melanin afﬁnity of metal ions. Acta Physiol. Scand., 104, 479–484. Hong, L., Liu, Y., and Simon, J. (2007) Binding of metal ions to melanin and their effects on aerobic reactivity. Photochem. Photobiol., 80, 477–411. Riley, P.A. (1997) Epidermal melanin: sunscreen or waste disposal? Biologist, 44, 408–411. Riley, P.A. (2010) The evolution of epidermal pigmentation: a speculative comment. Nederlands Tijsch. Derm. Vener., 20, 277–278. Borovanský, J. (1997) Detection of metals in tissues, cells and subcellular organelles. Sborník. Lek., 98, 77–97. Hong, L. and Simon, J.D. (2007) Current understanding of the binding sites, capacity, afﬁnity and biological signiﬁcance of metals in melanin. J. Phys. Chem. B, 111, 7938–7947. Eibl, O., Schultheiss, S., BlitgenHeinecke, P., and Schraermeyer, U. (2006) Quantitative chemical analysis of ocular melanosomes in the TEM. Micron, 37, 262–276. Potts, A.M. and Au, P.C. (1976) The afﬁnity of melanin for inorganic ions. Exp. Eye Res., 22, 487–491. White, L.P. (1958) Melanin: a naturally occurring cation exchange material. Nature, 158, 1427–1428. Pohla, H., Simonsberger, P., and Adam, H. (1983) X-ray microanalysis of rainbow trout (Salmo gairdneri, Rich) melanosomes with special reference to analytical methods. Mikroskopie, 40, 273–284. Borovanský, J., Hearn, R.R., Bleehen, S.S., and Russell, R.G.G. (1980) Distribution of 65Zn in mice with melanomas and in the subcellular

134

135

136

137

138

139

140

141

142

143

144

145

fractions of melanomas. Neoplasma, 26, 247–252. Bush, W.D. and Simon, J.D. (2007) Quantiﬁcation of Ca2+ binding to melanin supports the hypothesis that melanosomes serve a functional role in regulating calcium homeostasis. Pigment Cell Res., 20, 134–139. Borovanský, J. (1994) Zinc in pigmented cells and structures – interactions and possible roles. Sborník. Lék., 95, 309–320. Liu, Y., Hong, L., Kempf, V.E., Wakamatsu, K., Ito, S., and Simon, J.D. (2004) Ion exchange and adsorption of Fe(III) by Sepia melanin. Pigment Cell Res., 17, 262–269. Halliwell, B. and Gutteridge, J.M.C. (1999) Free Radicals in Biology and Medicine, 3rd edn, Oxford Science, Oxford, pp. 198–200. Szpoganicz, B., Gidanian, S., Kong, P., and Farmer, P. (2002) Metal binding by melanins: studies of colloidal dihydroxyindole-melanin, and its complexation by Cu(II) and Z(II) ions. J. Inorg. Biochem., 89, 45–53. Bogacz, A., Buszmann, E., and Wilczok, T. (1983) Competition between metal ions for DOPA melanin. Stud. Biophys., 132, 189–195. Borovanský, J. and Hach, P. (1999) Disparate behaviour of two melanosomal enzymes (α-mannosidase and γ-glutamyltransferase). Cell Mol. Biol., 45, 1047–1052. Bray, T.M. and Bettger, W.J. (1990) The physiological role of zinc as an antioxidant. Free Rad. Biol. Med., 8, 281–291. Willson, R.L. (1989) Zinc and iron in free radical pathology and cellular control, in Zinc in Human Biology (ed. C.F. Mills), Springer, Berlin, pp. 147–172. Borovanský, J. and Riley, P.A. (1989) Cytotoxicity of zinc in vitro. Chem. Biol. Interact., 69, 279–291. Borovanský, J., Horčičko, J., and Duchon, J. (1976) The hair melanosome: another tissue reservoir of zinc. Physiol. Bohemoslov., 25, 87–91. Borovanský, J., Blasko, M., Siracký, J., Schothorst, A.A., Smit, N.P.M., and

377

378

12 Physiological and Pathological Functions of Melanosomes

146

147 148

149

150

151

152

153

154

155

Pavel, S. (1997) Cytotoxic interactions of in vitro: melanoma cells are more susceptible than melanocytes. Melanoma Res., 7, 449–453. Gidanian, S., Mentelle, M., Meyskens, F.L., Jr, and Farmer, P.J. (2008) Melanosomal damage in normal human melanocytes induced by UVB and metal uptake – a basis for the pro-oxidant state of melanoma. Photochem. Photobiol., 84, 556–564. Pierpoint, C.G. (1983) Catecholate complexes. Coord. Chem. Rev., 38, 45–62. Hoogduijn, M.J., Smit, N.P., van der Laarse, A., van Niewpoort, A.F., Wood, J.M., and Thody, A.J. (2003) Melanin has a role in Ca++ homeostasis in human melanocytes. Pigment Cell Res., 16, 127–132. Panessa, B.J. and Zadunaisky, J.A. (1981) Pigment granules: a calcium reservoir in the vertebrate eye. Exp. Eye Res., 32, 593–604. Mattson, M.P. and Chan, S.L. (2003) Calcium orchestrates apoptosis. Nat. Cell Biol., 5, 1041–1043. Uemaetomari, I., Tabuchi, K., Nakamagoe, M., Tanaka, S., Murashita, H., and Hara, A. (2009) L-type voltagegated calcium channel is involved in the pathogenesis of acoustic injury in the cochlea. Tohoku J. Exp. Med., 218, 41–47. Barrenas, M.L. and Lindgren, F. (1991) The inﬂuence of eye colour on susceptibility to TTS in humans. Br. J. Audiol., 25, 303–307. Meyer zur Gottesberge, A. (1988) Physiology and pathophysiology of inner ear melanin. Pigment Cell Res., 1, 238–249. Gill, S.S. and Sait, A.N. (1997) Quantitative differences in endolymphatic calcium and endocochlear potential between pigmented and albino guinea pigs. Hear. Res., 113, 191–197. Lavado, A., Jeffery, G., Tovar, V., de la Villa, P., and Montoliu, L. (2006) Ectopic expression of tyrosine hydroxylase in the pigmented epithelium rescues the retinal abnormalities and visual function common in albinos in the absence of melanin. J. Neurochem., 96, 1201–1211.

156 Murillo-Cuesta, S., Contreras, J., Zurita,

157

158

159

160

161

162

163

164

165

166

E., Cediel, R., Cantero, M., Varela-Nieto, I., and Montoliu, L. (2009) Melanin precursors prevent premature agerelated and noise-induced hearing loss in albino mice. Pigment Cell Melanoma Res., 23, 72–83. Johnson, B.E., Mandel, G., Daniels, F., Jr (1972) Melanin and cellular reactions to ultraviolet radiation. Nature, 235, 147–148. MacDonald, C.J., Snell, R.S., and Lerner, A.B. (1965) The effect of laser radiation on the mammalian epidermal melanocyte. J. Invest. Dermatol., 45, 110–115. Momiyama, J., Hashimoto, T., Matsubara, A., Futai, K., Namba, A., and Shinkawa, H. (2006) Leupeptin, a calpain inhibitor, protects inner ear hair cells from aminoglycoside ototoxicity. Tohoku J. Exp. Med., 209, 89–97. Potts, A.M. (1962) The concentration of phenothiazines in the eye of experimental animals. Invest. Ophthalmol. Visual Sci., 1, 522–530. Larsson, B. (1998) The toxicology and pharmacology of melanin, in The Pigmentary System: Physiology and Pathophysiology (eds J.J. Nordlund, R.E., V.J. Hearing, R.A. King, W.S. Oetting, and J. Ortonne), Oxford University Press, New York, pp. 311–341. Lindquist, N.G. (1986) Melanin afﬁnity of xenobiotics. Uppsala J. Med. Sci., 91, 283–288. Miranda, M., Amicarelli, F., Bonﬁgli, A., Botti, D., Zarivi, O., and Poma, A. (1991) Changes of lipomelanosome membrane leakage versus pH, charge and composition. Melanoma Res., 1, 195–200. Cavatorta, P., Crippa, P.R., Ito, A.S., Casali, E., Ferrari, M.B., and Masotti, L. (1985) Fluorescence depolarization studies of melanosomal membranes from different sources. Physiol. Chem. Phys. NMR, 17, 365–370. Knörle, R., Schniz, E., and Feuerstein, T.J. (1998) Drug accumulation in melanin: an afﬁnity chromatographic study. J. Chromatogr. B, 714, 171–179. Gutmann, F. (1997) Charge Transfer Complexes in Biological Systems, Dekker, New York.

References 167 Tjalve, H., Nilsson, M., and Larsson, B.

168

169

170

171

172

173

174

175

176

(1981) Studies on the binding of chlorpromazine and chloroquine to melanin in vivo. Biochem. Pharmacol., 30, 1845–1847. Larsson, B., Oskarsson, A., and Tjalve, H. (1977) Binding of paraquat and diquat on melanin. Exp. Eye Res., 25, 353–359. Ito, A.S., Anzellini, G.C., Silva, S.C., Serra, O., and Szabo, A.G. (1992) Optical absorption and ﬂuorescence spectroscopy studies of ground state melanin–cationic porphyrin complexes. Biophys. Chem., 45, 79–89. Dencker, L., Larsson, B., Olander, K., Ullberg, S., and Yokota, M. (1979) False precursors of melanin as selective melanoma seekers. Br. J. Cancer, 39, 449–452. Larsson, H.S. (1991) Melanin-afﬁnic thioureas as selective melanoma seekers. Melanoma Res., 1, 85–90. Link, E.M., Brown, I., Carpenter, R.N., and Mitchell, J.S. (1989) Uptake and therapeutic effectiveness of 125I- and 211 At-methylene blue for pigmented melanoma in an animal model system. Cancer Res., 49, 4332–4337. Link, E.M. (1999) Targeting melanoma with 211At/131I-methylene blue: preclinical and clinical experience. Hybridoma, 18, 1877–1882. John, C., Bowen, W.D., Saga, T., Kinuya, S., Vilner, B.J., Baugold, J., Paik, C.H., Reba, R.C., Neumann, R.D., Varma, V.M., and McAfee, J.G. (1993) A malignant melanoma imaging agent: synthesis, characterization, in vitro binding and biodistribution of iodine125-(2-piperidinylaminoethyl)-4iodobenzamide. J. Nucl. Med., 34, 2169–2175. Guerquin-Kern, J.L., Hillion, F., Madelmont, J.C., Labarre, P., Papon, J., and Croisy, A. (2004) Ultrastructural cell distribution of the melanoma marker iodobenzamide: improved potentiality of SIMS imaging in life sciences. Biomed. Eng. Online, 3, 10–17. Labarre, P., Papon, J., Rose, A.H., Guerquin-Kern, J.L., Morandeau, L., Wu, T.D., Moreau, M.F., Bayle, M., Chezal, J.M., Croisy, A., Madelmont,

177

178

179

180

181

182

183

J.C., Turner, H., and Moins, N. (2008) Melanoma afﬁnity in mice and immunosuppressed sheep of [125I]N-(4dipropylaminobutyl)-4-iodobenzamide, a new targeting agent. Nucl. Med. Biol., 35, 783–791. Shan, L. (2009) Radiolabeled (Hetero) Aromatic Analogs of N-(2Diethylaminoethyl)-4-Iodobenzamide for Imaging and Therapy of Melanoma. Molecular Imaging and Contrast Agent Database (MICAD) [Internet], National Center for Biotechnology Information, Bethesda, MD. Leung, K. (2009) N-(2-(Diethylamino) Ethyl)-6-[18F]Fluoronicotinamide. Molecular Imaging and Contrast Agent Database (MICAD) [Internet], National Center for Biotechnology Information, Bethesda, MD. Chen, K.G., Valencia, J.C., Lai, B., Zhang, G., Paterson, J.K., Rouzaud, F., Berens, W., Wincovitch, S.M., Garﬁeld, S.H., Leapman, R.D., Hearing, V.J., and Gottesman, M.M. (2006) Melanosomal sequestration of cytotoxic drugs contributes to the intractability of malignant melanomas. Proc. Natl. Acad. Sci. USA, 103, 9903–9907. Gardette, M., Papon, J., Bonnet, M., Desbois, N., Labarre, P., Wu, T.D., Miot-Noirault, E., Madelmont, J.C., Guerquin-Kern, J.L., Chezal, J.M., and Moins, N. (2010) Evaluation of new iodinated acridine derivatives for targeted radionuclide therapy of melanoma using 125I, an Auger electron emitter. Invest. New Drugs, Epub ahead of print; doi: 10.1007/s10637-010-9471-x. Chen, K.G., Leapman, R.D., Zhang, G., Lai, B., Valencia, J.C., Cardarelli, C.O., Vieira, W.D., Hearing, V.J., and Gottesman, M.M. (2009) Inﬂuence of melanosome dynamics on melanoma drug sensitivity. J. Natl. Cancer Inst., 101, 1259–1271. Xie, T., Nguyen, T., Hupe, M., and Wei, M.L. (2009) Multidrug resistance decreases with mutations of melanosomal regulatory genes. Cancer Res., 69, 992–999. Stout, P.R. and Ruth, J.A. (1999) Deposition of [3H]cocaine, [3H ]nicotine and [3H]ﬂunitrazepam in mouse hair

379

380

12 Physiological and Pathological Functions of Melanosomes

184

185

186

187

188

189

190

191

192

melanosomes after systemic administration. Drug Metab. Dispos., 27, 731–735. Oxley, J.C., Smith, J.L., Kirschenbaum, L.J., and Marimganti, S. (2007) Accumulation of explosives in hair – part II: factors affecting sorption. J. Forensic Sci., 52, 1291–1296. Green, S.J. and Wilson, J.F. (1996) The effect of hair color on the incorporation of methadone into hair in the rat. J. Anal. Toxicol., 20, 121–123. Rollins, D.E., Wilkins, D.G., Krueger, G.G., Augsburger, M.P., Mizuno, A., O’Neal, C., Borges, C.R., and Slawson, M.H. (2003) The effect of hair color on the incorporation of codeine into human hair. J. Anal. Toxicol., 27, 545–551. Borges, C.R., Roberts, J.C., Wilkins, D.G., and Rollins, D.E. (2003) Cocaine, benzoylecgonine, amphetamine, and N-acetylamphetamine binding to melanin subtypes. J. Anal. Toxicol., 27, 125–134. Hayashibe, K., Mishima, Y., Ichihashi, M., and Kawai, M. (1986) Melanosomal antigenic expression on the cell surface and intracellular subunits within melanogenic compartments of pigment cells: analysis by antimelanosomeassociated monoclonal antibody. J. Invest. Dermatol., 87, 89–94. Sakai, C., Kawakami, Y., Law, L.W., Furumura, M., and Hearing, V.J., Jr (1997) Melanosomal proteins as melanoma-speciﬁc immune targets. Melanoma Res., 7, 83–95. Kawakami, Y., Robbins, P.F., Wang, R.F., Parkhurst, M., Kang, X., and Rosenberg, S.A. (1998) The use of melanosomal proteins in the immunotherapy of melanoma. J. Immunother., 21, 237–246. Marks, M.S., Theos, A.C., and Raposo, G. (2003) Melanosomes and MHC class II antigen-processing compartments. A tinted view of intracellular trafﬁcking and immunity. Immunol. Res., 27, 409–425. Rocha, N. and Neefjes, J. (2008) MHC class II molecules on the move for successful antigen presentation. EMBO J., 27, 1–5.

193 Palm, N..W. and Medzhitov, R. (2009)

194

195

196

197

198

199

200

Immunostimulatory activity of haptenated proteins. Proc. Natl. Acad. Sci. USA, 106, 4782–4787. Manini, P., Napolitano, A., Westerhof, W., Riley, P.A., and d’Ischia, M. (2009) A reactive ortho-quinone generated by tyrosinase-catalyzed oxidation of the skin-depigmenting agent monobenzone: self-coupling and thiol-conjugation reactions and possible implications for melanocyte toxicity. Chem. Res. Toxicol., 22, 1398–1405. Naish-Byﬁeld, S. and Riley, P.A. (1998) Tyrosinase autoactivation and the problem of the lag period. Pigment Cell Res., 11, 127–133. Kawakami, Y., Eliyahu, S., Sakaguchi, K., Robbins, P.F., Rivoltini, L., Yannelli, J.R., Appella, E., and Rosenberg, S.A. (1994) Identiﬁcation of the immunodominant peptides of the MART-1 human melanoma antigen recognized by the majority of HLA-A2restricted tumor inﬁltrating lymphocytes. J. Exp. Med., 180, 347–352. Douat-Casassus, C., Marchand-Geneste, N., Diez, E., Aznar, C., Picard, P., Geoffre, S., Huet, A., BourguetKondracki, M.L., Gervois, N., Jotereau, F., and Quideau, S. (2006) Covalent modiﬁcation of a melanoma-derived antigenic peptide with a natural quinone methide. Preliminary chemical, molecular modelling and immunological evaluation studies. Mol. Biosyst., 2, 240–249. Liu, J.B., Li, M., Chen, H., Zhong, S.Q., Yang, S., Du, W.D., Hao, J.H., Zhang, T.S., Zhang, X.J., and Zeegers, M.P. (2007) Association of vitiligo with HLA-A2: a meta-analysis. J. Eur. Acad. Dermatol. Venereol., 21, 205–213. Morgan, B.D.G., O’Neill, T., Dewey, D.L., Galpine, A.R., and Riley, P.A. (1981) Treatment of malignant melanoma by intravascular 4-hydroxyanisole. Clin. Oncol., 7, 227–234. Morgan, B.D.G. (1984) Recent results of a clinical pilot study of intra arterial 4HA chemotherapy in malignant melanoma, in Hydroxyanisole: Recent Advances in Anti-Melanoma Therapy (ed.

References P.A. Riley), IRL Press, Oxford, pp. 233–241. 201 Riley, P.A. (2003) Melanogenesis and melanoma. Pigment Cell Res., 16, 548–552. 202 Riley, P.A. (2004) Melanoma and the problem of malignancy. Tohoku J. Exp. Med., 204, 1–9. 203 Revskaya, E., Jongco, A.M., Sellers, R.S., Howell, R.C., Koba, W., Guimaraes, A.J., Nosanchuk, J.D., Casadevall, A., and Dadachova, E. (2009) Radioimmunotherapy of experimental

human metastatic melanoma with melanin-binding antibodies and in combination with dacarbazine. Clin. Cancer Res., 15, 2373–2379. 204 Dadachova, E. and Casadevall, A. (2005) Melanin as a potential target for radionuclide therapy of metastatic melanoma. Future Oncol., 1, 541–549. 205 Wood, J.M., Jimbow, K., Boissy, R.E., Slominski, A., Plonka, P.M., Slawinski, J., Wortsman, J., and Tosk, J. (1999) What’s the use of generating melanin? Exp. Dermatol., 8, 153–164.

381

wwwwwww

383

13 Dysplastic Nevi as Precursor Melanoma Lesions Stanislav Pavel, Nico P.M. Smit, and Karel Pizinger

13.1 Nevi as Risk Factors for Melanoma

Many scientists who are engaged in melanoma research are presently of the opinion that the development of cutaneous melanoma proceeds through various stages [1]. The basic idea is that some of the skin melanocytes can be transformed into benign pigmented nevus cells and certain nevus cells can eventually give rise to melanoma. The important change in the behavior of nevus cells is their reduced contact with keratinocytes, and the resulting limited ability to discharge the formed pigment and the tendency to clustering. Some of the pigmented nevi can later undergo more profound alterations, and develop macroscopically and microscopically visible dysplastic features. Only in exceptional cases, some of these dysplastic nevus cells may undergo malignant transformation. In 2002, a research group from Brisbane published a study investigating the risk factors for melanoma development in a young population (15–19 years) of melanoma patients [2]. The idea behind this research was that the crucial risk factors would manifest themselves at a younger age than the less important ones. The authors compared the data of 201 young melanoma patients with those of 196 healthy individuals matched for age, sex, and living area. The results convincingly showed that the most important melanoma risk factor in this group of young people was the number of pigmented nevi. Of lesser importance were factors such as light complexion (based on the susceptibility to sunburn), freckles, red hair, or melanoma in their family history. Surprisingly, there was no association between the development of melanoma and cumulative sun exposure. These results underline the importance of melanocytic nevi as a risk factor for melanoma development. 13.1.1 Development of Melanocytic Nevi

Melanocytic nevi start developing from the ﬁrst months of life. Clinical studies with twins [3] have provided evidence that the development of melanocytic nevi is Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

384

13 Dysplastic Nevi as Precursor Melanoma Lesions

under genetic control. In addition, the chance of the development of the acquired pigmented nevi is clearly related to skin pigmentation. Children with a light skin color run the highest risk of developing larger numbers of pigmented nevi [4]. The risk of development of dysplastic nevi is increased in people having large numbers of common acquired nevi and consequently the risk of developing dysplastic nevi is also greatest in lightly pigmented individuals [5]. In Japan, the incidence of dysplastic nevi is very low [6] and dysplastic nevi are unknown in people with very dark skin. 13.1.2 Description of Dysplastic Nevi

Dysplastic nevi were described for the ﬁrst time in the 1970s as large atypical pigmented moles that occurred especially in families with an increased incidence of melanoma [7]. While a large proportion of all common nevi disappear in elderly individuals by spontaneous regression, dysplastic nevi exhibit differing growth dynamics [8]. The clinical criteria that have been used in recent years for the recognition of these lesions are: (i) diameter 5 mm or greater, (ii) asymmetry, where one half of the lesion does not match the other with regard to color and shape, (iii) color variegation, (iv) border irregularity, and (v) evolution, when the nevus changes its appearance over time. Histological investigations of dysplastic nevi usually show a disrupted architecture and cytological atypia. The microscopic picture is reminiscent of the appearance of premalignant lesions such as those of the esophagus or cervix uteri.

13.2 Dysplastic Nevi as Precursor Lesions of Melanoma

The research of the last 30 years has provided clear evidence that pigmented nevi are associated with melanoma risk [9]. The strongest association has repeatedly been found in people with dysplastic nevi (see example in Figure 13.1). A metaanalysis of 46 investigations published before September 2002 showed that the presence of 101–120 common nevi (when compared with 15 nevi) was connected with a relative risk of 6.89 (95% conﬁdence interval 4.63, 10.25) [10]. In the population with dysplastic nevi, a similar risk factor was reached when patients had ﬁve (versus zero) dysplastic nevi. The large difference in the occurrence of dysplastic nevi in the normal population and in melanoma patients underlines the importance of dysplastic nevi as a signiﬁcant melanoma risk factor [11]. In the population with lightly pigmented skin, the presence of dysplastic nevi is usually found in less than 10% of individuals, whereas their presence in melanoma patients is signiﬁcantly higher (34–59%). Dysplastic nevi are also known to be direct precursors of melanoma (Figure 13.2). During the histopathological evaluation of excised melanomas, a high pro-

13.2 Dysplastic Nevi as Precursor Lesions of Melanoma a)

b)

nevus melanoma

Figure 13.1 (a) Photograph of the upper arm of a patient with multiple dysplastic nevi and one malignant melanoma arising from a nevus. (b) Detailed photograph of the nevus with superﬁcial melanoma (Breslow thickness 0.5 mm).

Photomicrograph of a vertical section of the skin showing a pigmented nevus with a malignant melanoma above it.

Figure 13.2

portion of melanoma tissue (up to 65%) contained remains of a nevoid component [12]. The malignant transformation of nevi can also be followed by dermatoscopy [13, 14]. Although the clinical and histological changes in dysplastic nevi are well recognized, knowledge of their speciﬁc biological and biochemical characteristics is limited. Some investigations have suggested that patients with multiple dysplastic

385

386

13 Dysplastic Nevi as Precursor Melanoma Lesions

nevi exhibit a DNA repair defect [15, 16]. However, this interesting association has not been further explored.

13.3 Cytological Differences between Normal Skin Melanocytes and Dysplastic Nevus Cells: Melanosomal and Mitochondrial Aberrations

As mentioned above, one of the characteristic features of nevus cells is their limited ability to transfer melanosomes to the surrounding keratinocytes. It is not clear what fundamental transfer mechanism is disturbed, but it leads to increased numbers of melanosomes in nevus cells [17]. The inability of nevus cells to discharge these melanin-producing organelles may increase the risk of development of intracellular oxidative imbalance. As mentioned elsewhere in this volume (see Chapter 12), melanosomes are a potential source of reactive oxygen species (ROS) and reactive o-dihydroxyphenol (catechol) derivatives that are potentially cyto- and genotoxic, and the structural integrity of melanosomes may play an important role in limiting this process. Several authors have observed the presence of large numbers of markedly aberrant melanosomes in dysplastic nevus cells when compared with common melanocytic nevi [17–21]. Rhodes et al. [20] reported that the percentage of abnormal melanosomes in dysplastic nevi was 7 times greater than that in common acquired melanocytic nevi and 22 times greater than that in normal skin melanocytes, but this percentage was only 20% less than in superﬁcial spreading melanoma. All the authors agree that the presence of melanosomal alterations can be considered as a useful marker of melanocytic atypia. The melanosomal abnormalities include not only structural defects of the melanosomal matrix, but also uneven and incomplete melanization. The melanization defects might be related to a higher leakage of reactive melanin precursors, which could lead to increased intracellular oxidative damage. From the studies dealing with the etiology of Parkinson’s disease it is well known that dopamine (one of the natural catechol derivatives) and its oxidation product (quinone) are toxic to mitochondria [22]. This toxicity may appear when vesicular storage of dopamine is disrupted. In vitro experiments showed that the addition of dopamine quinone to intact isolated mitochondria caused mitochondrial swelling that was completely prevented by cyclosporin A, suggesting opening of a permeability transition pore. Also, the addition of glutathione was able to block this quinone-induced mitochondrial swelling. Different molecular targets of dopamine quinone modiﬁcation in the mitochondrial proteome have already been characterized [23]. Important targets are tyrosyl residues in proteins. Their oxidative modiﬁcation to 3,4-dihydroxyphenylalanine (dopa) has recently been described in mitochondrial proteins, especially those with metal-binding properties [24]. This supports the idea of metal-catalyzed hydroxyl radical formation from mitochondrial superoxide and hydrogen peroxide. We can assume that a similar situation occurs in the pigment-producing cell with disrupted containment of melanogenesis and storage of melanin. Already

13.4 Metabolic Differences between Normal Skin Melanocytes and Dysplastic Nevus Cells

in 1988, Rhodes et al. [20] reported the presence of swollen mitochondria in dysplastic nevus cells, but this interesting observation apparently has not received much attention. One may expect that the presence of leaking melanogens (catechol derivatives) in the cytoplasm of dysplastic nevus cells, by virtue of their ability to produce ROS during redox-cycling reactions and their capability of liberating iron from ferritin (see Section 13.5), could damage mitochondria and cause mutations in mitochondrial DNA. This area of research could become critically important for a better understanding of the malignant transformation of pigment cells [25].

13.4 Metabolic Differences between Normal Skin Melanocytes and Dysplastic Nevus Cells: Preference for Pheomelanogenesis in Dysplastic Nevus Cells

As detailed in Chapters 4 and 6, pigment cells synthesize two types of pigment in their melanosomes: eumelanin and pheomelanin. The black/brown eumelanin polymer is able to efﬁciently absorb UV radiation and scavenge the ROS. The lighter pheomelanin polymer exhibits weaker UV absorption properties. This pigment can also generate ROS during UV exposure. In addition, during the synthesis of this pigment, the amino acid cysteine is consumed and that could lead to a decrease in the cellular concentration of glutathione – a crucially important intracellular antioxidant. We have recently shown that stimulation of melanogenesis in cultured melanocytes by increasing the tyrosine concentration leads to lower glutathione content [26]. The lowering of glutathione levels could have a wide range of harmful effects on cellular metabolism, including a pronounced retardation in DNA repair [27]. On the basis of their experiments, Land and Riley [28] concluded that the concentration of l-cysteine is an important determinant of the direction of melanogenesis. This is consistent with the so-called “three-step pathway” for mixed melanogenesis, as proposed by Ito and Wakamatsu [29]. According to this proposition, melanogenesis starts by the formation of cysteinyldopa and pheomelanin until the concentration of (utilized) cysteine is lowered. Then, eumelanogenesis starts to prevail. There are likely to be more factors involved in the switch between alternative pathways, including intramelanosomal transport of tyrosine and cysteine, the effect of local pH on the formation of dopaquinone, and other compounds of the pheo- and eumelanogenic pathways (see Chapter 6). Our own experiments with cultured human melanocytes have shown that when melanogenesis is stimulated, pigment cells isolated from lighter skin produce relatively more total pheomelanin and less total eumelanin than pigment cells originating from a darker skin [30]. This is in accordance with the in vivo situation. Lighter skin contains less melanin in general with a relatively higher percentage of pheomelanin. That makes the skin less resistant against the harmful effects of UV radiation and ROS.

387

388

13 Dysplastic Nevi as Precursor Melanoma Lesions

13.5 Pheomelanogenesis as a Possible Cause of Intracellular Oxidative Imbalance

The presence of oxygen is essential for the synthesis of melanin in the melanosomes. During this process, different reactive phenolic and indolic compounds are generated, and they polymerize with each other. However, some reactive molecules escape the polymerization process by leaking out of the melanosomes into the cytoplasm. Owing to their repeated oxidation followed by reduction (so-called redox cycling) these compounds form hydrogen peroxide, which can subsequently give rise to hydroxyl radicals. We have shown that these phenolic and indolic compounds are able to liberate iron from ferritin reserves [31], and that this situation can lead to oxidative DNA damage [32] (Figure 13.3). The presence of free iron cations in the cells can result in additional production of hydroxyl radicals due to the interaction of Fe(II) with hydrogen peroxide (the so-called Fenton reaction). Iron is able to bind tightly to melanin. The iron-binding sites are composed of catechol-like subunits [33]. These groups are present in high numbers in eumelanin structures, but not in the pheomelanin polymer. This means that the binding capacity of pheomelanin might not be sufﬁcient to bind the free iron liberated from ferritin reserves. The presence of a higher concentration of pheomelanin in the melanosomes would increase the risk of an increased intracellular concentration of free iron. If the overproduction of ROS is insufﬁciently compensated by

Simpliﬁed scheme of proposed pathways playing role in the hypermutability of dysplastic nevus cells.

Figure 13.3

13.7 Are Dysplastic Nevus Cells a Class of Cells Exhibiting a Mutator Phenotype?

various antioxidative defensive mechanisms this will result in oxidative stress. We have recently shown that melanosomes in dysplastic nevus cells contain signiﬁcantly more pheomelanin than normal melanocytes isolated from the same donors and the nevus cells exhibit clear presence of chronic oxidative stress [34]. As mentioned above, normal melanocytes partly lower the risk of oxidative stress by transferring melanosomes to surrounding keratinocytes. Dysplastic nevus cells, however, lose their contact with adjacent keratinocytes to a large extent. They cannot efﬁciently reduce their cellular load of melanosomes and this elevates the chance of oxidative stress with all the consequences. We have recently shown that DNA of dysplastic nevus cells contains more oxidative damage than DNA of normal melanocytes isolated from the same donors [26].

13.6 Dysplastic Nevus Cells as Senescent Cells

In recent years, some groups have described that cultured cells from common nevi grow for a short period, but at a certain point their cell division stops. This phenomenon is also clearly visible in cultured dysplastic nevus cells [26]. The growth termination is explained by the fact that the cells reach the stage of senescence. At that time, the cells acquire a senescent phenotype (i.e., their bodies become larger and the cells become thinner). Owing to their inability to divide, the cells cannot transfer possible mutations to the next generation. For this reason the senescent state is regarded as an essential barrier against malignant transformation [35]. Cellular senescence is also a programmed reaction to cell damage [36]. One example is senescence caused by a chronic stress situation. The regulation of senescence in melanocytes is not understood, but some researchers feel that the mechanism of senescence in melanocytes differs from the mechanisms in other cell types [37]. At any rate, it is clear that senescence of nevus cells does not guarantee complete protection. In some instances, senescence of these cells can be interrupted and (forced) cell division can cause the ﬁxation of mutations in the newly formed cells. It is not clear which factors play a role in the enforced cell division. It is possible that this factor could be ongoing oxidative damage in the pigment-producing cells.

13.7 Are Dysplastic Nevus Cells a Class of Cells Exhibiting a Mutator Phenotype?

DNA repair and DNA doubling in the course of cell division are processes that are under strict genetic control. That is why the frequency of DNA mutations in normal cells is low. This is, however, in sharp contrast with the high number of mutations found in cancer cells. The discrepancy in the number of mutations found in normal and cancer cells can be explained by the development of cells with a high mutation rate that can drive the carcinogenic process towards the generation of a malignant phenotype. The concept of a mutator phenotype in

389

390

13 Dysplastic Nevi as Precursor Melanoma Lesions

cancer was originally explained by mutations in DNA polymerases that cause them to become error-prone and/or mutations in DNA repair enzymes that diminish their ability to eradicate potentially mutagenic DNA lesions [38]. The mutator phenotype hypothesis has now been broadened to include factors involved in microsatellite instability, chromosomal instability, checkpoint instability, and maintenance of the epigenome [39]. Cells exhibiting this mutator phenotype have been described in various malignancies, such as prostate, gastrointestinal, and endometrial cancers [40, 41]. The opposing view is that the raised mutation rate is not itself the cause of tumor growth, but the consequence of clonal expansion combined with natural selection. It does not, however, fully exclude the importance of a raised mutation rate in certain stages of tumor development [42]. According to Traupe et al. [43], the mutation rate in multiple dysplastic nevi is exceptionally high and it can only be explained by a noninherited genetic defect (i.e., by a somatic mutation in the melanocytes). Such somatic mutations could give rise to a class of cell that occupies an intermediate stage between a normal melanocyte and a melanoma cell. As mentioned above, we have found that dysplastic nevus cells suffer from chronic oxidative stress and recent investigations by Dayal et al. [44] have shown that cells with the mutator phenotype can be maintained with increased intracellular production of hydrogen peroxide. Such a situation could be responsible for hypermutagenesis leading to genomic instability and the malignant phenotype. As dysplastic nevus cells fulﬁll these criteria, we propose that these cells could represent the mutator phenotype of pigmented cells that plays an essential role in the process of malignant transformation of melanocytes to malignant melanoma (Figure 13.4). However, the development of

Proposed scheme of the involvement of nevus cells exhibiting a mutator phenotype driving clonal evolution in the direction of malignant transformation.

Figure 13.4

References

melanoma from dysplastic nevus cells clearly requires the acquisition of malignant characteristics through further processes of mutation, cell selection, and clonal expansion. This is an exciting area for future research.

References 1 Hussein, M.R. (2005) Melanocytic

2

3

4

5

6

7

8

dysplastic naevi occupy the middle ground between benign melanocytic naevi and cutaneous malignant melanomas: emerging clues. J. Clin. Pathol., 58, 453–456. Youl, P., Aitken, J., Hayward, N., Hogg, D., Liu, L., Lassam, N., Martin, M., and Green, A. (2002) Melanoma in adolescents: a case-control study of risk factors in Queensland, Australia. Int. J. Cancer, 98, 92–98. Easton, D.F., Cox, G.M., Macdonald, A.M., and Ponder, B.A. (1991) Genetic susceptibility to naevi – a twin study. Br. J. Cancer, 64, 1164–1167. Gallagher, R.P., McLean, D.I., Yang, P., Coldman, A.J., Silver, H.K.B., Spinelli, J.J., and Beagrie, M. (1990) Suntan, sunburn and pigmentation factors and the frequency of acquired melanocytic nevi in children. Arch. Dermatol., 126, 770–778. Weinstock, M.A., Stryker, W.S., Stampfer, M.J., Lew, R.A., Willett, W.C., and Sober, A.J. (1991) Sunlight and dysplastic nevus risk. Results of a clinic-based case-control study. Cancer, 67, 1701–1706. Hara, K., Nitta, Y., and Ikeya, T. (1992) Dysplastic nevus syndrome among Japanese. A case study and review of the Japanese literature. Am. J. Dermatopathol., 14, 24–31. Clark, W.H.J., Reiemer, R.R., Greene, M.H., Ainsworth, A.A., and Mastrangelo, M.J. (1978) Origin of familial melanomas from heritable melanocytic lesions. “The B-K mole syndrome”. Arch. Dermatol., 114, 732–738. Halpern, A.C., Guerry, D., Elder, D.E., Trock, B., Synnestvedt, M., and Humpreys, T. (1993) Natural history of dysplastic nevi. J. Am. Acad. Dermatol., 29, 51–57.

9 Elder, D. (2010) Dysplastic naevi: an

update. Histopathology, 56, 112–120. 10 Gandini, S., Sera, F., Cataruzza, M.S.,

11

12

13

14

15

16

Pasquini, P., Abeni, D., Boyle, P., and Melchi, C.F. (2005) Meta-analysis of risk factors for cutaneous melanoma: I. Common and atypical naevi. Eur. J. Cancer, 41, 28–44. Friedman, R.J., Farber, M.J., Warycha, M.A., Papathasis, N., Miller, M.K., and Heilman, E.R. (2009) The “dysplastic” nevus. Clin. Dermatol., 27, 103–115. Sagebiel, R.W. (1993) Melanocytic nevi in histologic association with primary cutaneous melanoma of superﬁcial spreading and nodular types: effect of tumor thickness. J. Invest. Dermatol., 100, 322S–325S. Banky, J.P., Kelly, J.P., English, D.R., Yetman, J.M., and Dowling, J.P. (2005) Incidence of new and changed nevi and melanomas detected using baseline images and dermatoscopy in patients at high risk for melanoma. Arch. Dermatol., 141, 998–1006. Argenziano, G., Kittler, H., Ferrara, G., Rubegni, P., Malvehy, J., Puig, S., Cowell, L., Stanganelli, I., De Giorgi, V., Thomas, L., Bahadoran, P., Menzies, S.W., Piccolo, D., Marghoob, A.A., and Zalaudek, I. (2010) Slow-growing melanoma: a dermoscopy follow-up study. Br. J. Dermatol., 162, 267–273. Roth, M., Boyle, J.M., and Mulle, H. (1988) Thymine dimer repair in ﬁbroblasts of patients with dysplastic naevus syndrome (DNS). Experientia, 44, 169–171. Moriwaki, S.-I., Tarone, R.E., and Kramer, K.H. (1994) A potential laboratory test for dysplastic naevus syndrome: ultraviolet hypermutability of a shuttle vector plasmid. J. Invest. Dermatol., 103, 7–12.

391

392

13 Dysplastic Nevi as Precursor Melanoma Lesions 17 Vincente Ortega, V., Martinez Diaz, F.,

18

19

20

21

22

23

24

25

26

Carrascosa Romero, C., and Ortuno Pacheco, G. (1995) Abnormal melanosomes: ultrastructural markers of melanocytic atypia. Ultrastruct. Pathol., 19, 119–128. Takahashi, H., Horikoshi, T., and Jimbow, K. (1985) Fine structural characterization of melanosomes in dysplastic nevi. Cancer, 56, 111–123. Takahashi, H., Yamana, K., Maeda, K., Akutsu, Y., Horikoshi, T., and Jimbow, K. (1987) Dysplastic melanocytic nevus. Electron-microscopic observation as a diagnostic tool. Am. J. Dermatopathol., 9, 189–197. Rhodes, A.R., Seji, M., Fitzpatrick, T.B., and Stern, R.S. (1988) Melanosomal alterations in dysplastic nevi. A quantitative, ultrastructural investigation. Cancer, 61, 358–369. Beitner, H., Nakatani, T., and Hedblad, M.A. (1990) A transition electron microscopical study of dysplastic naevi. Acta. Derm. Venereol., 70, 411–416. Berman, S.B. and Hastings, T.G. (1999) Dopamine oxidation alters mitochondrial respiration and induces permeability transition in brain mitochondria: implication for Parkinson’s disease. J. Neurochem., 73, 1127–1137. Hasting, T.G. (2009) The role of dopamine oxidation in mitochondrial dysfunction: implications for Parkinson’s disease. J. Bioenerg. Biomembr., 41, 469–472. Zhang, X., Monroe, M.E., Chen, B., Chin, M.H., Heibeck, T.H., Schepmoes, A.A., Yang, F., Petritis, B.O., Camp, D.G., 2nd, Pounds, J.G., Jacobs, J.M., Smit, D.J., Bigelow, D.J., Smith, R.D., and Qian, W.J. (2010) Endogenous 3,4-dihydroxyphenylalanine and dopaquinone modiﬁcations on protein tyrosine: links to mitochondrially derived oxidative stress via hydroxyl radical. Mol. Cell. Proteomics, 9, 1199–1208. Czarnecka, A.M., Gammazza, A.M., Di Felice, V., Zummo, G., and Cappello, F. (2007) Cancer as a “mitochondriopathy”. J. Cancer Mol., 3, 71–79. Smit, N.P.M., Van Nieuwpoort, F.A., Marrot, L., Out, C., Poorthuis, B., Van Pelt, H., and Pavel, S. (2008) Increased

27

28

29

30

31

32

33

34

melanogenesis is a risk factor for oxidative DNA damage – study on cultured melanocytes and atypical nevus cells. J. Invest. Dermatol., 84, 550–555. Eiberger, W., Volkomer, B., Amouroux, R., Dhérin, C., Pablo Radicella, J., and Epe, B. (2008) Oxidative stress impairs the repair of oxidative DNA base modiﬁcations in human skin ﬁbroblasts and melanoma cells. DNA Repair, 7, 912–921. Land, E.J. and Riley, P.A. (2000) Spontaneous redox reaction of dopaquinone and the balance between the eumelanic and pheomelanic pathways. Pigment Cell Res., 13, 273–277. Ito, S. and Wakamatsu, K. (2008) Chemistry of mixed melanogenesis – pivotal role of dopaquinone. Photochem. Photobiol., 84, 582–592. van Nieuwpoort, F., Smit, N.P.M., Kolb, A.M., van der Meulen, J., Koerten, H.K., and Pavel, S. (2004) Tyrosine-induced melanogenesis shows differences in morphologic and melanogenic preferences of melanosomes from light and dark skin types. J. Invest. Dermatol., 122, 1251–1255. Pavel, S. and Smit, N.P.M. (1996) Detoxiﬁcation processes in pigmentproducing cells, in Melanogenesis and the Malignant Melanoma: Biochemistry, Cell Biology, Molecular Biology, Pathophysiology, Diagnosis and Treatment (eds Y. Hori, V.J. Hearing, and J. Nakayama), Elsevier, Amsterdam, pp. 161–168. Pavel, S., Smit, N.P.M., van der Meulen, J., Kolb, A.M., De Groot, A., Van der Velden, P.A., Gruis, N.A., and Bergman, W. (2003) Homozygous germline mutation of the CDKN2A/p16 gene and G-6-PD deﬁciency in a multiple melanoma case. Melanoma Res., 13, 171–178. Hong, L. and Simon, J.D. (2007) Current understanding of the binding sites, capacity, afﬁnity, and biological signiﬁcance of metals in melanin. J. Phys. Chem. B, 111, 7938–7947. Pavel, S., van Nieuwpoort, F., van der Meulen, J., Out, C., Pizinger, K., Cetkovská, P., Smit, N.P.M., and Koerten, H.K. (2004) Disturbed melanin synthesis

References

35

36

37

38

39

40

and chronic oxidative stress in melanoma precursor lesions. Eur. J. Cancer, 40, 1423–1430. Mooi, W.J. and Peeper, D.S. (2006) Oncogene-induced cell senescence – halting on the road to cancer. N. Engl. J. Med., 355, 1037–1046. Ben Porath, I. and Weinberg, R.A. (2005) The signals and pathways activating cellular senescence. Int. J. Biochem. Cell Biol., 37, 961–976. Ha, L., Merlino, G., and Sviderskaya, E.V. (2008) Melanomagenesis. Overcoming the barrier of melanocyte senescence. Cell Cycle, 7, 1944–1948. Loeb, L.A., Springgate, C.F., and Battula, N. (1974) Errors in DNA replications as a basis of malignant change. Cancer Res., 34, 2311–2321. Loeb, L.A., Bielas, J.H., and Beckman, R.A. (2008) Cancers exhibit a mutator phenotype: clinical implications. Cancer Res., 68, 3551–3561. Schwartz, S., Jr, Yamamota, H., Navarro, M., Maestro, M., and Perucho, M. (1999) Frameshift mutations at mononucleotide

41

42

43

44

repeats in caspase-5 and other target genes in endometrial and gastrointestinal cancer of the microsatellite mutator phenotype. Cancer Res., 59, 2995–3002. Colombo, P., Patriarca, C., Alfano, R.M., Cassani, B., Ceva Grimaldi, G., Roncalli, M., Bosari, S., Coggi, G., Campo, B., and Gould, V.E. (2001) Molecular disorders in transitional vs. peripheral zone prostate adenocarcinoma. Int. J. Cancer, 94, 383–389. Abdel-Rahman, W.M. (2008) Genomic instability and carcinogenesis: an update. Curr. Genomics, 9, 535–541. Traupe, H., Macher, E., Hamm, H., and Happle, R. (1989) Mutation rate estimates are not compatible with autosomal dominant inheritance of the dysplastic nevus “syndrome”. Am. J. Med. Genet., 32, 155–157. Dayal, D., Martin, S.M., Limoli, C.L., and Spitz, D.R. (2008) Hydrogen peroxide mediates the radiation-induced mutator phenotype in mammalian cells. Biochem. J., 413, 185–191.

393

wwwwwww

395

Index a N-acetyl-4-S-cysteaminylphenol (4-SCAP) 122, 132 N-acetylamphetamine 367 acid hydrolases 9, 10, 345 acid phosphatase 9, 10, 253, 257 actin 35, 46, 49, 269, 276, 295, 298, 301, 304, 306, 315 ADAMTS20 gene 327 adapalene 126, 142 adaptor protein (AP) complexes 270–273 – AP-1 272, 273 – AP-2 272 – AP-3 272, 274 – AP-3A 270 – AP-4 272 adaptor protein β3 334 adaptor protein δ1 334 adhesion molecules 313 adipose tissue, melanin in 37 adrenocorticotrophin (ACTH) 26, 47, 102, 148 age-related macular degeneration (AMD) 204, 214–216 age spots 333 agouti signaling protein (ASIP) 26, 103, 329, 331 albinism 95, 102, 194, 216, 248, 251, 253, 271 albino cochleas 32 albino mammals 32 all-trans-retinoic acid (ATRA) 121–123, 126, 141 aloesin 123, 125, 135 α-hydroxy acids (AHAs) 142 amphetamine 367 amphibian melanophores 28, 296–298 antibiosis 63 antioxidant properties 201, 350,351

antioxidant protein 1 330 antioxidants 36, 127, 136–138, 213 AP3D1 gene 99, 121, 128, 138, 145, 208, 237–290, 328, 334 apoptosis 363 arbutin 120, 123, 132, 133, 135 α-arbutin 122, 132 arylsulfatase 9 ascorbic acid 124, 125, 136, 137 ASP gene 329 asparagine-linked glycosylation 128 ATOX1 gene 330 ATP7A gene 253, 330 ATP7B gene 330 ATPase 9, 107, 257, 266, 302 ATPase 7α 330 ATPase 7β 330 atrial arrhythmia 35 ATRN gene 330 azelaic acid 123, 125, 134

b baboon substantia nigra 228 barnacles, adhesion of 64 BCL2 gene 328 bee venom 148 benzamide 366 benzothiazolecarboxylic acid (BTCA) 178, 179 benzoylecgonine 367 bergapten (5-MOP) 148 Berzelius, J.J. 4 biogenesis of lysosome-related organelle complexes (BLOC) 273–276 – BLOC-1 270, 273, 274 – BLOC-2 270, 275 – BLOC-3 270, 275, 276 Birbeck, M.S.C. 1 bisisolylmaleimide GF 109203X 126

Melanins and Melanosomes: Biosynthesis, Biogenesis, Physiological, and Pathological Functions, First Edition. Edited by Jan Borovanský and Patrick A. Riley. © 2011 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2011 by Wiley-VCH Verlag GmbH & Co. KGaA.

396

Index black human hair 168 – photoemission electron microscopy analysis of 196 Blaschko, H. 1 blue irides 199 BMY-28565 128 bone morphogenetic proteins (BMPs) 39, 146 Bowman−Birk protease inhibitor 141 brain – biological role of neuromelanin in the human brain 232, 233 – melanocytes 36, 37 – nonclassical murine melanocytes 45 brown irides 199 N-butyldeoxynojircimicin 123, 128 4-n-butylresorcinol 122, 125, 132

c C2-ceramide 122, 127 cadherins 49, 313 calcium 25, 32, 34, 36, 49, 128, 216, 217, 363, 364 calcium d-pantethein-S-sulfonate 128 cAMP 147 β-carotene 126, 142 carotenoids 202 caspase-3 238 catalase 139 cataract surgery 200 catechol oxidases 64, 81, 92, 94, 98 catechols 66, 69, 70, 80, 82, 98, 130, 351–353, 363 cathepsin B 10 cathepsin D 197, 217 cathepsin L 10 Caucasian eye 188 cellular reductants, oxidation of 210–212 centaureidin 140 cephalopods 168 ceramide glucosyltransferase 279 Chediak−Higashi syndrome (CHS) 248, 269, 333, 334 chemical peels 142 choroid 30, 44, 187, 250 – melanin content in the 189, 193 chromatoblasts 45 chromatophores 23, 24, 45 – from donkey conjuctiva 3 ciglitazone 148 ciliary body 30, 44, 187 cisplatin 366, 367 classical melanocytes 22, 23, 38–41 – distribution and function of 25–28

– genes involved in development of 39, 40 – melanosome transfer from 47–51 – transfer of melanin from 45–51 – see also nonclassical melanocytes clathrin 255 coat-colour dilution 251 cocaine 367 cochlea 31–33, 51 – albino cochleas 32 competitive inhibitors 130 complex I 237, 238 copper 79–82, 92–99, 119, 134, 206, 332 cornea 201 CTNS gene 330 cuticular hardening 65 cyanophores 24 cyclodopa (leucodopachrome) 70, 76–78, 89, 169, 188 cys-dopamine melanin (DA-M) 239–241 cystamine 124, 138 cysteamine 138 cysteine 24, 71, 77, 118, 169, 188, 237, 303, 332, 353 cysteinyldopa 36, 77, 118, 188, 189 5-S-cysteinyldopa 71, 176, 353, 355 6-cysteinyldopa 71 5-S-cysteinyldopaquinone 77 cystinosin 330 cytochrome c 238 cytokines 127, 145, 146 cytophagocytosis 309, 310 cytoskeleton, molecular motors and the 248, 276, 277

d Darier’s disease 313 dark skin, distribution of melanin granules 313 DCT gene 335 DEFB103A gene 330 β-defensin 3 330 defensins 26 deoxyarbutin 123, 125, 133 deoxymannojirimycin 123, 128 depigmenting agents 121–143 – acting prior to melanin synthesis 121–129 – classiﬁcation of 122–124 – clinical applications of 125, 126 – transcriptional inhibition of melanogenic enzymes 121–128 dermis, melanocytes in the 27, 28 DHI-2-carboxylic acid (DHICA) 72, 90, 136, 170, 188, 252, 346

Index – oligomers formed by oxidation of 173 – oxidation of 172 cis-diaminedichloroplatinum II, see cisplatin dickkopf1 (DKK1) 28, 329, 332 dihydrolipoic acid 122, 127, 128 1,4-dihydroxybenzene 131 5,6-dihydroxyindole (DHI) 72, 90, 169, 171, 188, 252, 346 – oligomers formed by oxidation of 172 3,4-dihydroxyphenylalanine (dopa) 118, 169, 188, 355, 356 N,N´´-dilinoleylcystamine 124, 139 2,6-dimethoxy-N-(4-methoxyphenyl) benzamide 123 diosgenin 122, 127 disodium isostearyl 2-O-lascorbyl phosphate 124, 137 DKK1 gene 329 docosahexaenoic acid 198 dolichol 232 dopa 64, 169, 188, 355, 356 L-dopa-positive melanocytes 34 dopachrome 21, 73, 76, 136, 188, 345 dopachrome tautomerase (Dct) 21, 87, 98–100, 104, 136, 170, 188, 252, 335 dopamine 227, 232 dopamine melanin (DA-M) 239–241 dopaquinone 21, 76, 77, 88, 118, 169, 188 , 255, 346, 387 – addition to the terminal amino group of a lysine residue in a protein chain 347 – electrophilic addition to a cysteine residue in a protein chain 347 dynactin 300, 301, 314 dynein(s) 298–301, 314, 315 dynesin(s) 295 dysplastic nevi – cytological differences between normal skin melanocytes and 386, 387 – description of 384 – hypermutability of 388 – metabolic differences between normal skin melanocytes and 387 – mutator phenotype 389–391 – as precursor lesions of melanoma 384–386 – as senescent cells 389

e E-cadherin 49, 313 ear – acoustic injury 363 – cochlea 31–33 – melanocytes of the inner ear

31–33

– sound/heat conversion 360 – vestibular labyrinth 33 early endosomal antigen 1 (EEA1) 255 EDN1 gene 326, 328, 329 EDN3 gene 326, 328, 329 EDNRB gene 40, 326, 328, 329 EGFR gene 327 ellagic acid 123, 125, 135 endocytic pathway 263–265 – organelles 254–256 endolymph 31 endoplasmic reticulum (ER) 265, 266 ȕ-endorphin 27 endosomal−melanosomal network 268 endosomal sorting complex required for transport (ESCRT) 255, 266 endosomal tubules 268 endothelin 1 329 endothelin 3 329 endothelin B receptor 329 enzymes – melanogenic 92–102 – in melanosomes 9 epidermal melanin unit 7 epidermis – melanocytes in the 25 – renewal of 141 epithelial cells 33, 187 erythrophores 24 etoposide 367 eumelanin(s) 21, 24, 25, 37, 78, 143, 167, 168, 189, 232, 343, 387 – analysis of 180 – biosynthesis of 89 – degradative studies 176–178 – eumelanin/pheomelanin switch 78, 170 – oxidative degradation of 177 – structure and properties of 174 eumelanogenesis 74, 78, 102, 103, 129, 143, 171, 387 eumelanosomes 22 – biogenesis of 248 – in dark irides 200 – stages 249 exocytosis 310, 311 eye 188, 247 – color regulation 330–332 – cornea 201 – lens 200 – melanin content in the choroid 193 – melanin content in the iris 193, 194 – melanin content in the pigmented structures of the 190–194 – melanocytes of the 28, 29

397

398

Index

f fatty acids 105, 129 ferritin 128, 233, 388 fetal human RPE cells, apoptosis of 208, 209 FGF gene 326 FGFR2 gene 326 ﬁbroblast growth factors 43 ﬁbroblasts 27, 28 ﬁlopodia 296, 311, 312, 315 ﬁlopodial-phagocytosis model 311, 312 ﬁsh melanophores 28, 296–298 ﬁsh, RPE cells 299 Fitzpatrick, Thomas B. 1, 3, 10 ﬂavonoids 134, 135 ﬂuorescent emission 358 follicular lymphoma 328 follicular melanin unit 10 forkhead box D3 (FoxD3) 39, 329 forskolin 147 FOXD3 gene 326, 328, 329 Foxn1 48, 314 frog melanophores 297, 298 FZD4 gene 326

g G-protein-coupled receptor (GPCR) GPR143 252 β-galactosidase 9 gallic acid 124 genes 323 – and diseases involved with regulating melanocyte function 329, 330 – involved in classical melanocyte development 39, 40 – involved in development of RPE 43 – involved in distribution of melanosomes 337, 338 – involved in melanin production 334–336 – involved in melanoblast development, migration and speciﬁcation 324–327 – involved in melanocyte differentiation, survival and proliferation 325–328 – involved in melanosome movement 336, 337 – involved in regulating melanocyte function 327–333 – involved in the biogenesis of melanosomes and other lysosome-related organelles 333, 334 – involved in transfer of melanosomes 337 geniposide 145 gentisic acid 123, 133, 134

gentisic acid methyl ester 123 glabridin 143 glucosamine 123 β-glucuronidase 9 γ-glutamyltransferase 9 glutathione 32, 92, 131, 138, 237, 353, 386 glutathione S-transferase (GST) α4 (Gsta4) gene 32 S-glutathionylation 237, 238, 241 glutathionyldopa 355 glycolic acid 126, 142 glycoproteins 49, 50 glycosphingolipids 279 GM-CSF gene 329 GNA11 gene 328 GNAQ gene 328 gnetol 123, 135 gp100, see Pmel17 gp100/silver protein 330, 335 GPNMB gene 335 GPR143 gene 335 granulocyte macrophage colony-stimulating factor 329 Grb2 145 Griscelli syndrome 269, 295, 337, 337 Griscelli syndrome 1 (GS1) 302 Griscelli syndrome 2 (GS2) 303 growth factors 145, 146 GTPases 255, 269 guanine nucleotide exchange factor (GEF) 302 gunmetal (Rab geranylgeranyl transferase α subunit) 334

h hair – black human hair 168 – color regulation 330–332 – isolation of melanosomes from 7 – melanocytes in the follicle 25 – PEEM analysis of black hair 196 – PEEM analysis red hair 196 – pheomelanin in red hair 179 Harderian gland 30, 31 Harding-Passey melanoma melanosomes 349 heart, melanocytes 33–36, 44, 45 hemocyanins 92 hemophagocytic lymphohistiocytosis 303 hemophagocytic syndrome 303 hepatocyte growth factor receptor 329 Hermansky−Pudlak syndrome (HPS) 248, 269, 271–276, 333, 334

Index – adaptor protein complexes 271–273 – BLOC complexes 273–276 HGF gene 327 Hirschsprung’s disease 325 Hirschsprung’s disease type 2 326, 328 HMB45 265 homotypic fusion and vacuole protein sorting (HOPS) complex 270, 277, 278 HPS1 gene 334 HPS2 gene 334 HPS3 gene 334 HPS4 gene 334 HPS5 gene 334 HPS6 gene 334 HPS7 gene 334 HPS8 gene 334 human eye 188 human fundus 188 human leukocyte antigen A2 (HLA-A2) 369 human red hair pheomelanin 179 human RPE cell line, effects of exogenous melanosomes on 208 human substantia nigra 228, 229 hydrogen bonding 348, 349 hydrogen peroxide 139, 352 hydroquinone 122, 125, 131 hydroquinone monobenzyl ether 122 6-hydroxy-3,4-dihydrocumarins 124, 138 p-hydroxy-methoxybenzene (4-hydroxyanisole) 122, 132, 357, 369 4-hydroxyanisole (p-hydroxymethoxybenzene) 122, 132, 357, 369 4-hydroxyphenyl α-D-glucopyranoside (α-arbutin) 132 hydroxystilbene compounds 135 hyperpigmentation 117, 332, 333 hypopigmentation 117, 129, 141, 332 hypoxia-inducible factor (HIF) 99

i IFNG gene 330 IL1 gene 330 inﬂammation 26 inhibitor strength 130 insects – cuticular hardening in 65 – defensive secretions 64 interferon-γ 330 interfollicular melanocytes 26, 27 interleukin-1 (IL-1) 330 interleukin-6 (IL-6) 127 intracellular melanosome transport 299–307 intracellular oxidative imbalance 388, 389 ionization thresholds 196

iridophores 24 iris 30, 44, 187 – blue irides 199 – brown irides 199 – darkening of the 194 – eumelanosomes in dark irides 200 – melanin content in the 193, 194 – role as a ﬁlter of light 198–200 iris pigment epithelial (IPE) cells 193 iron 37, 99, 139, 206–208, 233, 235, 361, 362, 388 isotretinoin 142 ITGB1 gene 327

j Jimbow, Kowichi

8

k keratinocyte growth factor (KGF) 49, 337 keratinocyte growth factor receptor (KGFR) 312 keratinocytes 7, 25, 45–51, 117 – fate of melanin in 313–315 – melanosome positioning within 51 KGF gene 337 KIF13A protein 270, 276 kinectin 300 kinesin-2 298 kinesins 295, 299–301, 315 KIT gene 326, 328, 329 Kit oncogene 329 KIT/SCF gene 326, 328 KITL gene 329, 332 kojic acid 123, 125, 133 Kunitz-type trypsin inhibitor 141

l latanoprost 194 leaky melanosomes 353–355 lectins 49, 50, 313 LEF1 gene 326 lens, aging of 200 Lepidium apetalum extract 122, 127 leucodopachrome (cyclodopa) 89, 169, 188 leucophores 24 Lewy bodies 234, 235, 239 Lewy neurites 239 licorice extracts 126, 143 light scattering 358 light skin, distribution of melanin granules 313 linoleic acid 105, 123, 126, 129, 142, 143 lipids 147, 148, 279 – in ocular melanosomes 197, 198 – peroxidation by semiquinone radical 353

399

400

Index lipoamide dehydrogenase 138 lipocalin-type prostaglandin D synthase (L-PGDS) 27, 36 lipofuscin 192, 208, 225, 229 α-lipoic acid (LA), see thioctic acid liquiritin 143 locus coeruleus 227, 229 lysophosphatidic acid (LPA) 122, 126 lysosomal hydrolases 9, 247, 256 lysosomal membrane proteins (LAMP) 1 and 2 253 lysosomal trafﬁcking regulator (Lyst) 269 lysosome-related organelles (LROs) 10, 247, 256–258, 33, 248, 256–259, 333 lysosomes 10, 197, 250, 256–258 – enzymes 253 LYST gene 334

m macromelanosomes 263 macules, white 332 magnesium L-ascorbate-3-phosphate 124, 137 mahogunin ring ﬁnger 1 330 malignant melanoma, aberrant melanosomes in 354, 368 mammalian melanocytes 298, 299 mammals, albino mammals 32 Manchester Pulse Radiolysis apparatus 76 α-mannosidase 10 MART-1 251, 266, 335, 369 MART-1 tumor-associated antigen 253 MATP protein (SLC45A2) 251, 253, 330, 332, 335 matrix proteins 8, 347–349 MCIR gene 329 MCOLN3 gene 327 medulla oblongata 227, 229 melagenin 146 Melan-A cells 129 melanin granules 2, 5, 30, 194, 214 melanin(s) 2, 21 – in adipose tissue 37 – agents acting during synthesis of 129–139 – antioxidant properties of 201–209 – biochemistry 118, 119 – biosynthesis 118 – biosynthesis of 87–116 – biosynthetic studies 169–176 – broadband absorption spectra 198 – classiﬁcation and general properties of 168, 169 – content in melanosomes 345

– content in pigmented structures of the eye 190–194 – content in the choroid 193 – content in the iris 193, 194 – content in the retinal pigment epithelium 190–193 – degradative studies 176–179 – development of chemistry of 11 – exploitation of proteins as speciﬁc targets in melanoma therapy 368, 369 – fate in the keratocyte 313–315 – free radical nature of 350–352 – genes involved in production of 334–336 – molecular and cellular transfer mechanisms 47–50 – ocular 187–189 – role in light transmission through the RPE and choroid 200, 201 – roles of 26 – structure of 167–185 – synthesis in mammals 323 – transfer from classical and nonclassical melanocytes 45–51 – transfer from nonclassical melanocytes 51 melano-phagolysosomes 314 melano-phagosomes 314 melanoblasts 21, 34, 38 – genes involved in development, migration and speciﬁcation of 324, 325, 326, 327 – late determined 40, 41 melanocortin-1 receptor (MC1R) 25, 26, 102, 103, 144, 331 melanocortin-2 receptor 329 melanocortins 102 melanocyte-directed enzyme prodrug therapy 358 melanocyte-stimulating hormone (MSH) 26 α-melanocyte-stimulating hormone (α-MSH) 26, 27, 37, 102, 104, 119, 144, 306 melanocytes 11, 21, 22, 24, 132, 187, 324 – of the brain and neuromelanins 36, 37 – classical 21, 22 – dendrite formation inhibition 140 – in the dermis 27, 28 – development of nevi 383, 384 – in the epidermis 25 – of the eye 28, 29, 42–44 – factors and signaling pathways regulating function of 331 – genes and diseases involved with regulating 329, 330 – genes involved in differentiation, survival and proliferation of 325–327, 328

Index – genes involved in regulating function of 327–333 – in the hair follicle 25 – of the heart 33–36, 44, 45 – of the inner ear 31–33 – interfollicular 26, 27 – intracellular trafﬁcking proteins 270, 271 – mammalian 298, 299 – marker of 27 – of the murine eye 42–44 – of the murine heart 44, 45 – nonclassical 21, 22 – of the RPE 42, 43 – uveal melanocytes 43, 44 – see also classical melanocytes; nonclassical melanocytes melanogenesis 87–116 – beneﬁts of 65 – control of 78 – early stages 169–171 – enhancers 143–148 – epidermal 118 – inhibitors and enhancers of 117–165 – interference with the melanogenic pathway 138, 139 – mixed melanogenesis 175, 176 – nonenzymatic formation of intermediates 74–77 – outline of reactions 75 – paracrine signaling and regulation of epidermal 119, 120 – phase I 78, 79 – radicals and reactive species associated with 352–355 – Raper−Mason pathway 88–92 – regulation of 102–107 – role of o-quinones in 74–82 – study methods 120 melanogenesis enhancers 143–148 – cytokines and growth factors 145, 146 – forskolin and cAMP 147 – lipids 147, 148 – melanotropic peptides 144, 145 – oligonucleotides and p53 activation 147 – piperin 147 melanogenic cells – deﬁnition 21–24 – distribution and function of 24–37 – embryonic development of 37, 38 melanogenic enzymes, post-translational modiﬁcation of 128, 129 melanolipofuscin 190, 192, 205, 214 melanolysomes 190, 192

melanoma 50, 197, 199, 200, 257, 259, 349, 353 – detection or treatment 365, 366 – dysplastic nevi as precursion lesions of 384–386 – nevi as risk factor for 383, 384 – seekers 365 – therapy 368–369 melanoma antigen 1 335 melanoma cells 7 melanoma melanosomes 7 melanophilin (Mlph) 269, 299, 303, 304, 337 melanophores 24 – from ﬁsh and amphibians 296–298 – in lower vertebrates 28 melanoproteins 2, 11 melanoregulin 337 melanosomal lipids 7, 197–198 melanosomal proteins 8, 9, 196, 197, 251–253 melanosome degradation 10 melanosome maturation 269 melanosome research – biochemical 4 – history of 1–20 – in the post-Seiji era 9–11 – in the pre-Seiji era 1–5 – in the Seiji era 5–9 melanosome shape 195–196, 349–350 melanosome staging 5,22, 249–250, 258, 261 melanosome terminology 5–-6 melanosome transfer 46–51, 309–313 – from classical melanocytes 47–51 – cytophagocytosis 309 – exocytosis 310, 311 – ﬁlopodial-phagocytosis model 311, 312 – inhibitors of 139–141 – reduction of 140 melanosome transport 46, 295–308 melanosomes 5, 21, 45, 46, 187, 249–253, 295–322 – actin-based transport 301–307 – afﬁnity for polycyclic and other compounds 196, 364–368 – announcement of independent status in Medical News 2 – antioxidant properties of 201–209, 350–352 – biochemical studies 7–9 – biogenesis of 247–294 – as centers of free radical activity 350–357 – chemical composition 196–198, 344–346

401

402

Index – – – – – – –

components 251–253, 267, 268, 344–346 concentration 344–345 as dark structures in migmatic rocks 12 deﬁnition 6, 11, 12 developmental stages 5 disintegration and degradation 10 distribution of metabolites in defective 354 – distribution of metabolites in normal 354 – early-stage 263–265 – endocytic system and formation of 254–268 – as energy transducers 358–360 – and the ER 265, 266 – exploitation of proteins as speciﬁc targets in melanoma therapy 368, 369 – fate of 7 – formation in eye pigment cells 250 – function of ﬁbrillar melanosomes 262 – functional microanatomy of 346–350 – genes involved in distribution of 337, 338 – genes involved in movement of 336, 337 – genes involved in the biogenesis of 333, 334 – genes involved in the transfer of 337 – genetics of structure and function of 323–341 – intercellular transfer of 296 – intracellular transport 299–307 – intracellular transport of 300 – isolation of 4, 8, 10 – long-term deposition of compounds in 367, 368 – as lysosome-related organelles (LROs) 256–259, 333 – maturation stages 5, 6, 22 – and metal ions 8, 206, 360–364 – microanatomy of 196, 346 – microtubule-based transport 300, 301 – motility in RPE 307, 308 – mutation 269–279 – number of entries in the ISI Web of Knowledge 11, 12 – ocular 187–189 – ontogenesis of 5 – organization of 261, 346 – origin of 263–268 – participation in chemoresistance 366, 367 – photochemical reactions 360 – photoionization threshold of 351 – photon/phonon conversion 359, 360 – positioning within keratinocytes 51

– pro-oxidant effects of interactions with metal ions 213, 214 – Rab27a-Mlph(melanophilin)-Myo5a tripartite protein complex 304–306 – segregation from endocytic pathway beyond stage I melanosomes 266, 267 – sound/heat conversion 360 – as speciﬁc targets in diagnosis and therapy 365–369 – stage I 5, 249, 259 – stage II 5, 250, 252, 259 – stage III 5, 250, 259, 267 – stage IV 5, 250, 259, 267 – studies showing presence of enzymes in 9 – terminology 5–9 – as a therapeutic target 356, 357 – tissue concentration of 343, 344 – transport in mouse RPE cells 308 – transport of 46, 295–308 – ultrastructural and histochemical studies 6, 7 melanotropic peptides 144, 145 melasma 333 membrane-associated transporter protein (MATP) 251, 253, 330, 332, 335 membrane glycosylation, inhibition of 141 Menkes disease 253 MET gene 327, 329 metal ions, melanosomes and 360–364 metals 8, 37, 66, 134, 206, 216, 360 – sequestration of 206–207, 361 methimazole 124, 125, 139, 365 5-methoxypsoralen (5-MOP) 148 8-methoxypsoralen (8-MOP) 148 methylene blue (MTB) 365, 366 methylophiopogonanone B 140 MGRN1 gene 330 microphthalmia 329 microphthalmia-associated transcription factor (MITF) 40, 43, 103, 104, 119, 121, 306, 307 microtubules 46, 269, 276, 298, 300, 301 MITF gene 326, 328, 329 MITF-M 27, 39, 40, 119, 126 mitochondria, neuromelanin effects on function of 237–239 MLPH gene 337 MNT-1 cells 249, 268 mocha mice 272 monobenzone 357 monoclonal antibodies 368, 369 mouse – melanocytes 297 – RPE cells 299

Index MREG gene 337 multivesicular bodies (MVBs) 255, 256, 280 mussels 64 MUTED gene 334 MYO5A gene 337 Myo5a-GT 306 MYO7A gene 337 myosin Ib 271, 276, 277 myosin Va (Myo5a) 269, 295, 298, 299, 301, 302, 315, 337 myosin VIIa (Myo7a) 299, 337 MyoX 312 Myrip (Slac2-c) 307

n neoglycoproteins 313 neural crest cells (NCC) 34, 37–41, 44, 45 neurogulin 1 329 neuromelanin granules 226 neuromelanin-mediated cell death 237 neuromelanin-pigmented neuron, from the human substantia nigra 226 neuromelanin(s) 36, 37, 225–246 – biological role in the human brain 232, 233 – catabolism pathways 231 – comparison of the cytoxicity of neuromelanin with synthetic DA-M 239–241 – comparison of the effects of neuromelanin and DA-M in vitro and in vivo 240 – cytotoxicity mechanisms 235–237 – description of 225–227 – development and metabolism of 227–231 – dopamine-based 232 – effects on mitochondrial function 237–239 – effects on the UPS 239 – free radical scavenging abilities 232, 233 – importance in Parkinson’s disease 225–246 – and increase with age 227 – involvement in neurological disease 233–235 – as a metal binder 233 – norepinephrine-based 232 – phylogenetic development of 227 – protein degradation 233 – structure of 231, 232 – in substantia nigra 4, 234 – in vitro and in vivo effects 235–241 nevi, as risk factors for melanoma 383, 384 niacinamide 125, 140, 313 nicotinamide 366

nitric oxide 202 NLRP1 gene 328 nocodazole 308 nonclassical melanocytes 22, 23, 28–37, 41, 42 – melanocytes of the eye 28, 29 – transfer of melanin from 45–51 – see also classical melanocytes norepinephrine 227 NRG1 gene 329, 332

o OA1/GPR143 251 OA1 protein 252, 253 – and melanosome biogenesis 263 obesity 37 ocular albinism (OA) 248, 335 ocular melanin – antioxidant properties of 201 – biogenesis of 187–189 – role as a broadband optical ﬁlter 198–201 – testing protective effects in cultured cells 207, 208 ocular melanosomes – molecular composition of 196–198 – morphology of 195, 196 – pro-oxidant properties of 209–216 – proteins 196, 197 – roles of 217 – structure of 194–198 oculocutaneous albinism (OCA) 248, 335 oculocutaneous albinism (OCA) type 1 252, 335 oculocutaneous albinism (OCA) type 2 252, 335 oculocutaneous albinism (OCA) type 3 335, 336 oculocutaneous albinism (OCA) type 4 253, 335 oligonucleotides 147 optical density, of choroid melanin 200 osteopetrosis 335 OSTM1 gene 335 oxyresveratrol 123, 135 oxyresveratrol derivative 123

p P-cadherin 49, 313 P gene 329, 335 P locus 332 P protein (OCA2) 102, 251, 252, 329, 335 p38 mitogen-activated protein kinase (MAPK) 105 p53 activation 147 PAH gene 335

403

404

Index paired box gene 6 329 pallidin 334 palmitic acid 105, 129 D-pantetheine-S-sulfonate 123 PAR2 gene 51, 337 Parkinson’s disease 37, 231, 234, 386 – substantia nigra 234 PAX3 gene 39, 326 PAX6 gene 326, 329 pearl mice 272 peroxidase inhibitors 139 peroxisome proliferator-activated receptor (PPAR) activators 144, 148 peroxygenase 139 peroxynitrite 202 pH 189, 257 phagocytosis 309 phenolic thioether amines 357 phenoloxidases 92 phenothiazine derivatives 364 phenylalanine hydroxylase 335 phenylketonuria 335 pheomelanin(s) 21, 26, 36, 78, 167–169, 189, 232, 343, 387 – analysis of 180 – biosynthesis of 91, 92 – degradative studies 178, 179 – main degradation products of 178 – production of 171 pheomelanogenesis 78, 91, 92, 103 – in dysplastic nevus cells 387 – late stages of 174, 175 – as a possible cause of intracellular oxidative imbalance 388, 389 phospholipase A2 148 phospholipase D2 123 photochemical reactions 360 photodegradation 205 photoemission electron microscopy (PEEM) 196, 198, 203 photon/phonon conversion 359, 360 photoprotection 26, 28, 31, 46, 66, 247, 324 photoreception 28 photosensitizing agents 148 – quenching of electronically excited states of 203–206 phototherapeutic drugs 204 phytoncide 124 piceatonnol 124 piebaldism 325, 328 pigment dispersion syndrome 335 pigment transport – model systems to study 296–299 – in RPE cells 299

pigmentation 65, 66 – regulation of 104–107 piperin 147 PLDN gene 334 Pmel17 101, 250–252, 258, 261, 264, 277, 330, 332, 335, 350 – amlyoid ﬁbrils 262 – biosynthesis and amyloid formation 260–262 – and ﬁbrillar matrix for melanin deposits 259, 260 – and generation of early-stage melanosomes 259–262 – structure of 259 Pmel17-Mα 262 poikilotherms 359 polycyclic compounds 364–368 polyunsaturated lipids, oxidation of 203 POMC gene 329 postinﬂammatory hyperpigmentation 333 premelanosome ﬁbrils, model for formation of 261 premelanosomes 5, 6, 187, 249 prodrugs 134, 357, 358 proopiomelanocortin 329 N-propionyl-4-S-cysteaminylphenol 357 prostaglandin E2 147 prostaglandins 147, 148, 194 protease-activated receptor-2 (PAR-2) 48, 49, 312, 313, 337 – inhibition of 140, 141 protein-bound dopa 355, 356 protein haptenation 368, 369 protein hydroperoxides 355 protein kinase A (PKA) 119 protein kinase C inhibitors 122 proteinase-activated receptor 2 (PAR-2), see protease-activated receptor-2 (PAR-2) proteins, in ocular melanosomes 196, 197 proteomic analysis of melanosomes 345–346 proximal Raper−Mason pathway 88–90 psoralens 148 psoralens plus UV-A (PUVA) 148 pterorhodin 24 pulse radiolysis 74–77 – rate constants 77 pycnogenol 124, 125 pyridinium bisretinoid (A2E) 205 pyrrole-2,3-dicarboxylic acid (PDCA) 177, 178 pyrrole-2,3,5-tricarboxylic acid (PTCA) 177, 178, 180 pyrrole carboxylic acids 177 pyrroloquinoline 123

Index

q quinolines 123 quinone(s) 63, 123, 352, 353 – redox potentials 69 o-quinones 356, 357, 368, 370 – addition−elimination (substitution) reactions 73 – addition reactions 71 – antibiosis 63, 64 – balanid adhesion 64 – biological chemistry of 63–86 – cuticular hardening in insects 65 – cyclization of 72, 73 – cycloaddition 67, 68 – defensive secretions 64 – general biological signiﬁcance of 63–66 – intramolecular addition 72, 73 – pigmentation 65, 66 – polymerization 71, 72 – polymerization of 71, 72 – reactivity 66–73 – reduction 68–70 – role in melanogenesis 74–82 – structure and reactivity 66–68 p-quinones 63 Quintox mechanism 80, 82

r Rab geranylgeranyl transferase 302 Rab geranylgeranyl transferase II 270, 278 Rab GTPases 50, 278, 311 Rab3GEF 302 Rab4 255 Rab5 255 Rab7 270, 279, 301, 330 Rab11 255, 278, 279 Rab27a 269, 278, 299, 302, 303, 306, 307, 337 Rab27a-binding domain (R27BD) 306 Rab27a-Mlph(melanophilin)-Myo5a tripartite protein complex 295, 299, 304–308, 315 Rab27a-Myrip-Myo7a (myosin VIIa) 295 Rab32 270, 278 Rab38 270, 278 Rab38 gene 334 RABGGTA gene 334 radicals associated with melanogenesis 352–355 Raper−Mason pathway 75, 88–92, 170, 352 Ras-associated protein 7 330 Ras-associated protein 27a 337 Ras-associated protein 38 334 rat substantia nigra 228

RB1 328 reactive oxygen species (ROS) 26, 131, 136, 201, 237, 352, 386 – generation of 210–212 red hair – pheomelanin 179 – photoemission electron microscopy analysis of 196 redox-active metal ions, sequestration of 206, 207 redox exchange 69, 70, 169 redox potentials, of quinones 69 reducing agents 136–138 resveratrol 123, 127, 128, 135 retinal aging 214–216 retinal pigment epithelium (RPE) 22, 28, 29, 44, 187–189, 195, 250, 258, 295, 299, 307, 308 – aging of 215 – effects of exogenous melanosomes on 208 – genes involved in development of 43 – lipids 197 – melanin content in the 189, 190–193 – melanin granules, cytotoxic properties of aged 214–216 – melanocytes 42, 43 – melanosomes, aging of 217 retinitis pigmentosum 327, 328 retinoblastoma 328 retinoids 141, 142 riboﬂavin 208

s salicylic acid 126, 142 s.c.CAT-DISORT model 343 Seiji, Makoto 1, 3, 9 SEMA3C gene 328 SEMA4A gene 328 semiquinone species 232, 352 senescence 389 Sepia ink 168 sepia melanin 178 serine protease inhibitors 312 sialylated O-linked oligosaccharides 260 SILV, see Pmel17 silver protein 101 Simon, Gustav 1 singlet oxygen 203–206 skin – color regulation 330–332 – distribution of melanin granules 313 skin-lightening agents 131 skin pigmentation 315 SLC7A11 330

405

406

Index SLC24A5 251, 253, 330, 332 SLC25A5 335 SLC45A2 251, 253, 330, 332, 335 Slp2-a 304 SNA12 gene 326 SNAP25 (25-kDa synaptosome-associated protein) 311 solar lentigo 121 soluble N-ethylmaleimide-sensitive factor attachment protein receptors (SNAREs) 50, 248, 277, 311 solute carrier 7 A11 330 solute carrier 24 A5 330, 335 solute carrier 45 a2 330 sound/heat conversion 360 SOX2 gene 329 SOX9 gene 326, 329 SOX10 gene 326 SOX10 gene 329 SOX18 gene 329 sphingolipids 147, 148 sphingomyelin 126 Sry box gene 2 329 Sry box gene 9 329 Sry box gene 10 329 Sry box gene 18 329 standardized testing of pigmentary regulators (STOPR) 120 stem cell factor 329 stromal melanocytes 193 substantia nigra 225–229 – neuromelanin in 4 – neuromelanin-pigmented neurons from human at various ages 230 superoxide ions 352 superoxide radical 213 synaptobrevin, see vesicle-associated membrane protein (VAMP) synaptotagmin-like protein (Slp) 303 syntaxin 311 syntaxin-13 277, 278 α-synuclein 235

t tanning responses 332, 333 tarazotene 126, 141, 142 target SNAREs 277 TCFAP2A gene 326 terminology of melanosomes 5 4-tertiary butylcathecol 122, 132 4-tertiary butylphenol 122, 131, 132 tetanus-insensitive vesicle-associated membrane protein (TI-VAMP) 277 thioctic acid 124, 125, 127, 138

thiols 71, 89, 138, 237, 356 thiouracil 365 thiourea 365 thioureylenes 365 tissue concentration of melanosomes 343, 344 TNFA gene 330 α-tocopherol 124, 125, 137 α-tocopherol ferulate 124, 137 tocopherols 138 transforming growth factor (TGF)-β1 127 trichloroacetic acid 126 4,5´, 8-trimethylpsoralen 148 trioxsalen 148 TRPM1 253 TRPM7 253 tryptophan-2,3-dioxygenase 9 TSG101 (ESCRT-1) 271 tumor necrosis factor 330 tumor necrosis factor-α 127 tunicamycin 123 TYR gene 329, 335 tyrosinase 21, 87, 88, 92, 170, 188, 229, 252, 279, 329, 335, 369 – catalytic cycle of 95–98 – degradation of 105–106 – catalytic site 119 – catecholase (dopa oxidase) reaction cycle 98 – cresolase (tyrosine hydroxylase) reaction cycle 96–98 – function of 251 – maturation and degradation 118 – melanin synthesis and 129–136 – regulation of levels of 104–106 – structure of 92–95 – ubiquination 129 tyrosinase activation 78, 79 tyrosinase inactivation 79–82 tyrosinase-related protein 1 (Tyrp1) 21, 87, 100, 101, 118, 170, 251, 252, 275, 279, 329, 335, 336 tyrosinase-related protein 2 (Tyrp2), see dopachrome tautomerase (Dct) tyrosinase-speciﬁc activity, control of 106, 107 tyrosine hydroxylase 32 tyrosine-2-oxoglutarate amino transferase 9 L-tyrosine, to L-dopachrome 88–90 TYRP1 gene 329, 335

u ubiquitin−proteasome system (UPS) 105, 239 uncompetitive inhibitor 130

Index Usher syndrome type 1B 307, 337 UV radiation 25, 26, 47, 103, 145, 171, 247, 314, 328, 332 UV responses 333 uveal melanocytes 28, 29, 30, 43, 44 – choroid 44 – ciliary body 44 – iris 44

vinblastine 367 visible pigmentation 328 Visudyne® 204 vitamin D analogs 146 vitamin E 137 vitiligo 117, 132, 146, 327, 328, 332, 357 Vps33 278 VPS33A gene 334

v vacuolar protein sorting 33A 334 VAMP, see vesicle-associated membrane protein (VAMP) Vax proteins 43 vertebrates, classical and nonclassical melanocytes in 21–61 verteporﬁn 204 vesicle-associated membrane protein (VAMP) 311 vesicle-associated membrane protein (VAMP7) 277 vesicle SNAREs 277 vesicles 267 vestibular inner ear melanocytes (VIEMs) 33

w Waardenburg−Shah syndrome 328 Waardenburg syndrome 325, 328 WNT1 gene 326 WNT3A gene 326 Wnts 39

x xanthophores 24 xenobiotics 204, 216

z zeolites 122, 127 zinc 5, 93, 99, 216, 303, 362, 363

407