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The Pigmentary System: Physiology and Pathophysiology

The Pigmentary System: Physiology and Pathophysiology Edited by

James J. Nordlund Dermatologist and Professor Emeritus, Group Health Associates, Cincinnati, OH, USA

Raymond E. Boissy Professor of Dermatology and Cell Biology, University of Cincinnati College of Medicine, Cincinnati, OH, USA

Vincent J. Hearing Chief, Pigment Cell Biology Section, Laboratory of Cell Biology, National Institutes of Health, Bethesda, MD, USA

Richard A. King Director, Genetics Division, Department of Medicine, Institute of Human Genetics, University of Minnesota, Minneapolis, MN, USA

William S. Oetting Associate Professor, Genetics Division, Department of Medicine Institute of Human Genetics, University of Minnesota, Minneapolis, MN, USA

Jean-Paul Ortonne Professor of Dermatology and Chairman, Department of Dermatology, University of Nice-Sophia Antipolis, Nice, France

SECOND EDITION

© 2006 Blackwell Publishing Ltd Blackwell Publishing, Inc., 350 Main Street, Malden, Massachusetts 02148-5020, USA Blackwell Publishing Ltd, 9600 Garsington Road, Oxford OX4 2DQ, UK Blackwell Publishing Asia Pty Ltd, 550 Swanston Street, Carlton, Victoria 3053, Australia The right of the Author to be identiﬁed as the Author of this Work has been asserted in accordance with the Copyright, Designs and Patents Act 1988. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by the UK Copyright, Designs and Patents Act 1988, without the prior permission of the publisher. First published 1998 by Oxford University Press Second edition 2006 1 2006 Library of Congress Cataloging-in-Publication Data The pigmentary system : physiology and pathophysiology / edited by James J. Nordlund . . . [et al.]. — 2nd ed. p. ; cm. Includes bibliographical references and index. 1. Pigmentation disorders. 2. Melanocytes. I. Nordlund, James J. [DNLM: 1. Pigmentation Disorders–physiopathology. 2. Melanocytes. 3. Pigmentation–physiology. WR 265 P6309 2006] RL790.P53 2006 616.5¢5–dc22 2005030369 A catalogue record for this title is available from the British Library Set in 9/12 pt Sabon by SNP Best-set Typesetter Ltd., Hong Kong Printed and bound in India by Replika Press PVT Ltd ISBN-13: 978-1-4051-2034-0 ISBN-10: 1-4051-2034-7 Commissioning Editor: Stuart Taylor Editorial Assistant: Saskia Van der Linden Development Editor: Rob Blundell Production Controller: Kate Charman For further information on Blackwell Publishing, visit our website: http://www.blackwellpublishing.com The publisher’s policy is to use permanent paper from mills that operate a sustainable forestry policy, and which has been manufactured from pulp processed using acid-free and elementary chlorine-free practices. Furthermore, the publisher ensures that the text paper and cover board used have met acceptable environmental accreditation standards. Blackwell Publishing makes no representation, express or implied, that the drug dosages in this book are correct. Readers must therefore always check that any product mentioned in this publication is used in accordance with the prescribing information prepared by the manufacturers. The author and the publishers do not accept responsibility or legal liability for any errors in the text or for the misuse or misapplication of material in this book.

Contents

Contributors, x Foreword, xv Preface, xvii Acknowledgements, xviii Frontispiece can be found between pages ii and iii Part I: The Physiology of the Pigmentary System Section 1: Historical and Comparative Perspectives of the Pigmentary System, 3 1 A History of the Science of Pigmentation, 5 Sidney N. Klaus 2 Comparative Anatomy and Physiology of Pigment Cells in Nonmammalian Tissues, 11 Joseph T. Bagnara & Jiro Matsumoto Section 2: The Science of Pigmentation, 61 3 General Biology of Mammalian Pigmentation, 63 Walter C. Quevedo Jr. & Thomas J. Holstein 4 Extracutaneous Melanocytes, 91 Raymond E. Boissy & Thomas J. Hornyak 5 Regulation of Melanoblast Migration and Differentiation, 108 David M. Parichy, Mark V. Reedy, & Carol A. Erickson 6 Melanoblast Development and Associated Disorders, 140 Richard A. Spritz 7 Biogenesis of Melanosomes, 155 Raymond E. Boissy, Marjan Huizing, & William A. Gahl 8 Melanosome Trafﬁcking and Transfer, 171 Glynis A. Scott 9 Melanosome Processing in Keratinocytes, 181 H. Randolph Byers 10 The Regulation of Melanin Formation, 191 Vincent J. Hearing 11 The Tyrosinase Gene Family, 213 William S. Oetting & Vijayasaradhi Setaluri 12 Molecular Regulation of Melanin Formation: Melanosome Transporter Proteins, 230 Murray H. Brilliant 13 Transcriptional Regulation of Melanocyte Function, 242 Kazuhisa Takeda & Shigeki Shibahara 14 Enzymology of Melanin Formation, 261 Francisco Solano & José C. García-Borrón

15 Chemistry of Melanins, 282 Shosuke Ito & Kazumasa Wakamatsu 16 The Physical Properties of Melanins, 311 Tadeusz Sarna & Harold A. Swartz 17 Photobiology of Melanins, 342 Antony R. Young 18 Toxicological Aspects of Melanin and Melanogenesis, 354 Edward J. Land, Christopher A. Ramsden, & Patrick A. Riley 19 Regulation of Pigment Type Switching by Agouti, Melanocortin Signaling, Attractin, and Mahoganoid, 395 Gregory S. Barsh 20 Human Pigmentation: Its Regulation by Ultraviolet Light and by Endocrine, Paracrine, and Autocrine Factors, 410 Zalfa Abdel-Malek & Ana Luisa Kadekaro 21 Paracrine Interactions of Melanocytes in Pigmentary Disorders, 421 Genji Imokawa 22 Growth Factor Receptors and Signal Transduction Regulating the Proliferation and Differentiation of Melanocytes, 445 Ruth Halaban & Gisela Moellmann 23 Aging and Senescence of Melanocytes, 464 Debdutta Bandyopadhyay & Estela E. Medrano 24 The Genetics of Melanoma, 472 Vanessa C. Gray-Schopfer & Dorothy C. Bennett 25 The Transformed Phenotype of Melanocytes, 489 Dong Fang & Meenhard Herlyn Part II: The Pathophysiology of Pigmentary Disorders Section 3: An Overview of Human Skin Color and its Disorders, 497 26 A More Precise Lexicon for Pigmentation, Pigmentary Disorders, and “Chromatic” Abnormalities, 499 James J. Nordlund, Tania Cestari, Pearl Grimes, Henry Chan, & Jean-Paul Ortonne 27 The Normal Color of Human Skin, 504 James J. Nordlund & Jean-Paul Ortonne 28 Mechanisms that Cause Abnormal Skin Color, 521 Jean-Paul Ortonne & James J. Nordlund v

CONTENTS

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vi

Section 4: Disorders of Hypopigmentation, Depigmentation and Hypochromia, 539 Genetic Hypomelanoses: Disorders Characterized by Congenital White Spotting — Piebaldism, Waardenburg Syndrome, and Related Genetic Disorders of Melanocyte Development — Clinical Aspects, 541 Richard A. Spritz Genetic Hypomelanoses: Acquired Depigmentation, 551 Rozycki Syndrome, 551 Jean L. Bolognia Vitiligo Vulgaris, 551 James J. Nordlund, Jean-Paul Ortonne, & I. Caroline Le Poole Genetic Hypomelanoses: Generalized Hypopigmentation, 599 Oculocutaneous Albinism, 599 Richard A. King & William S. Oetting Albinoid Disorders, 613 Philippe Bahadoran & Jean-Paul Ortonne Ataxia Telangiectasia, 621 Anne-Sophie Gadenne Hallerman–Streiff Syndrome, 623 James J. Nordlund Histidinemia, 623 Marnie D. Titsch Homocystinuria, 625 Allan D. Mineroff Oculocerebral Syndrome with Hypopigmentation, 626 Jean L. Bolognia Tietz Syndrome, 630 Jean-Paul Ortonne Kappa-Chain Deﬁciency, 631 Jean-Paul Ortonne Menkes’ Kinky Hair Syndrome, 631 Tanusin Ploysangam Phenylketonuria, 634 Allan D. Mineroff Genetic Hypomelanoses: Localized Hypopigmentation, 636 “Hypomelanosis of Ito” and Mosaicism, 636 Wolfgang Küster & Rudolf Happle Focal Dermal Hypoplasia, 645 James J. Nordlund Hypomelanosis with Punctate Keratosis of the Palms and Soles, 646 Jean L. Bolognia Darier–White Disease (Keratosis Follicularis; 124200), 647 Jean L. Bolognia Nevus Depigmentosus, 649 Stella D. Calobrisi Tuberous Sclerosis Complex, 652 Pranav B. Sheth Genetic Hypomelanoses: Disorders Characterized by Hypopigmentation of Hair, 657

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39

Bird-Headed Dwarﬁsm (Seckel Syndrome), 657 Stan P. Hill Down Syndrome, 658 Rosemary Geary Fisch Syndrome, 659 Stan P. Hill Premature Canities, 659 James J. Nordlund Mandibulofacial Dysostosis (Treacher Collins Syndrome), 660 Rosemary Geary Myotonic Dystrophy, 660 Peggy Tong PHC Syndrome (Böök Syndrome), 661 Stan P. Hill Pierre Robin Syndrome, 661 James J. Nordlund Prolidase Deﬁciency, 662 Pranav B. Sheth Metabolic, Nutritional, and Endocrine Disorders, 664 Metabolic and Nutritional Hypomelanoses, 664 Peter S. Friedmann Hypomelanosis Associated with Endocrine Disorders, 667 Peter S. Friedmann Chemical, Pharmacologic, and Physical Agents Causing Hypomelanoses, 669 Chemical and Pharmacologic Agents Causing Hypomelanoses, 669 Stefania Briganti, Monica Ottaviani, & Mauro Picardo Physical Agents Causing Hypomelanoses, 683 Jean-Philippe Lacour Infectious Hypomelanoses, 686 Jean-Philippe Lacour Inﬂammatory Hypomelanoses, 699 Jean-Philippe Lacour Hypomelanoses Associated with Melanocytic Neoplasia, 705 Lieve Brochez, Barbara Boone, & Jean-Marie Naeyaert Miscellaneous Hypomelanoses: Depigmentation, 725 Alezzandrini Syndrome, 725 Wiete Westerhof, David Njoo, & Henk E. Menke Idiopathic Guttate Hypomelanosis, 726 Wiete Westerhof, David Njoo, & Henk E. Menke Leukoderma Punctata, 729 Wiete Westerhof, David Njoo, & Henk E. Menke Lichen Sclerosus et Atrophicus, 731 Philippe Bahadoran Vagabond Leukomelanoderma, 732 Wiete Westerhof, David Njoo, & Henk E. Menke Vogt–Koyanagi–Harada Syndrome, 734 Wiete Westerhof, David Njoo, & Henk E. Menke Westerhof Syndrome, 741 Wiete Westerhof, David Njoo, & Henk E. Menke

CONTENTS

40 Miscellaneous Hypomelanoses: Hypopigmentation, 745 Disseminated Hypopigmented Keratoses, 745 Wiete Westerhof, David Njoo, & Henk E. Menke Hypermelanocytic Punctata et Guttata Hypomelanosis, 746 Wiete Westerhof, David Njoo, & Henk E. Menke Progressive Macular Hypomelanosis, 748 Henk E. Menke, Germaine Relyveld, David Njoo, & Wiete Westerhof Sarcoidosis, 751 Wiete Westerhof, David Njoo, & Henk E. Menke 41 Miscellaneous Hypomelanoses: Extracutaneous Loss of Pigmentation, 754 Alopecia Areata, 754 Wiete Westerhof Heterochromia Irides, 756 Wiete Westerhof, David Njoo, & Henk E. Menke Senile Canities, 760 Wiete Westerhof, David Njoo, & Henk E. Menke Sudden Whitening of Hair, 764 Wiete Westerhof, David Njoo, & Henk E. Menke 42 Hypochromia without Hypomelanosis, 767 Jean-Philippe Lacour Section 5: Disorders of Hyperpigmentation and Hyperchromia, 769 43 Genetic Epidermal Syndromes: Disorders Characterized by Generalized Hyperpigmentation, 771 Adrenoleukodystrophy, 771 Sheila S. Galbraith & Nancy Burton Esterly Familial Progressive Hyperpigmentation, 774 Nancy Burton Esterly, Eulalia Baselga, Beth A. Drolet, Susan Bayliss Mallory, & Sharon A. Foley Fanconi Anemia, 776 Sheila S. Galbraith & Nancy Burton Esterly Gaucher Disease, 778 Sheila S. Galbraith & Nancy Burton Esterly 44 Genetic Epidermal Syndromes: Disorders Characterized by Reticulated Hyperpigmentation, 780 Berlin Syndrome, 780 Eulalia Baselga & Nancy Burton Esterly Cantu Syndrome, 781 Eulalia Baselga & Nancy Burton Esterly Kindler Syndrome, 781 Eulalia Baselga & Nancy Burton Esterly Dermatopathia Pigmentosa Reticularis, 784 Eulalia Baselga & Nancy Burton Esterly Dyschromatosis Universalis Hereditaria, 786 Eulalia Baselga & Nancy Burton Esterly Epidermolysis Bullosa with Mottled Pigmentation, 788 Eulalia Baselga & Nancy Burton Esterly Familial Mandibuloacral Dysplasia, 790 Eulalia Baselga & Nancy Burton Esterly Hereditary Acrokeratotic Poikiloderma, 792 Eulalia Baselga & Nancy Burton Esterly

Hereditary Sclerosing Poikiloderma, 795 Eulalia Baselga & Nancy Burton Esterly Mendes Da Costa Disease, 796 Eulalia Baselga & Nancy Burton Esterly Naegeli–Franceschetti–Jadassohn Syndrome, 798 Eulalia Baselga & Nancy Burton Esterly Reticulated Acropigmentation of Dohi (Dyschromatosis Symmetrica Hereditaria), 799 Eulalia Baselga & Nancy Burton Esterly Reticulate Acropigmentation of Kitamura, 802 Eulalia Baselga & Nancy Burton Esterly Rothmund–Thomson Syndrome, 804 Eulalia Baselga & Nancy Burton Esterly 45 Genetic Epidermal Syndromes with Café-au-lait Macules, 809 Familial Multiple Café-au-lait Spots, 809 Nancy Burton Esterly Neuroﬁbromatosis, 809 Nancy Burton Esterly, Eulalia Baselga, & Sheila S. Galbraith Neuroﬁbromatosis 1 with Noonan Syndrome, 816 Nancy Burton Esterly McCune–Albright Syndrome, 817 Sheila S. Galbraith & Nancy Burton Esterly Segmental Neuroﬁbromatosis, 819 Nancy Burton Esterly & Eulalia Baselga Silver–Russell Syndrome, 820 Nancy Burton Esterly, Eulalia Baselga, & Sheila S. Galbraith Watson Syndrome, 823 Nancy Burton Esterly 46 Genetic Epidermal Pigmentation with Lentigines, 824 Lentigo Simplex, 824 Mary K. Cullen Lentigo Senilis et Actinicus, 829 Mary K. Cullen Centrofacial Lentiginosis, 837 Mary K. Cullen LEOPARD Syndrome, 842 Mary K. Cullen Carney Complex, 851 Mary K. Cullen Other Lentiginoses, 863 Mary K. Cullen 47 Genetic Epidermal Syndromes: Localized Hyperpigmentation, 873 Anonychia with Flexural Pigmentation, 873 Nancy Burton Esterly, Eulalia Baselga, & Beth A. Drolet Incontinentia Pigmenti, 873 Sheila S. Galbraith & Nancy Burton Esterly Periorbital Hyperpigmentation, 879 Nancy Burton Esterly, Eulalia Baselga, & Beth A. Drolet vii

CONTENTS

Pigmentary Demarcation Lines, 880 Sheila S. Galbraith & Nancy Burton Esterly Dowling–Degos Disease, 882 Sheila S. Galbraith & Nancy Burton Esterly 48 Genetic Epidermal Syndromes: Disorders of Aging, 884 Acrogeria, 884 Nancy Burton Esterly, Eulalia Baselga, Peter M. H. Chan, & Beth A. Drolet Metageria, 886 Nancy Burton Esterly, Eulalia Baselga, Peter M. H. Chan, & Beth A. Drolet Progeria, 886 Nancy Burton Esterly, Eulalia Baselga, Peter M. H. Chan, & Beth A. Drolet Xeroderma Pigmentosum, 889 Anita P. Sheth, Nancy Burton Esterly, Eulalia Baselga, Peter M. H. Chan, & Beth A. Drolet Werner Syndrome, 894 Nancy Burton Esterly, Eulalia Baselga, Peter M. H. Chan, Beth A. Drolet, & Cindy L. Lamerson 49 Congenital Epidermal Hypermelanoses, 898 Dyskeratosis Congenita, 898 Susan Bayliss Mallory, Peggy L. Chern, & Sharon A. Foley Ectodermal Dysplasias, 901 Susan Bayliss Mallory, Peggy L. Chern, & Sharon A. Foley Transient Neonatal Pustular Melanosis, 905 Susan Bayliss Mallory, Peggy L. Chern, & Sharon A. Foley Universal Acquired Melanosis, 906 Susan Bayliss Mallory, Peggy L. Chern, & Sharon A. Foley 50 Acquired Epidermal Hypermelanoses, 907 Acanthosis Nigricans, 907 Norman Levine & Cynthia Burk Acromelanosis Progressiva, 914 Norman Levine & Cynthia Burk Becker Nevus, 915 Norman Levine & Cynthia Burk Café-au-lait Spots, 917 Norman Levine & Cynthia Burk Carcinoid Syndrome, 919 Norman Levine & Cynthia Burk Conﬂuent and Reticulated Papillomatosis, 922 Norman Levine & Cynthia Burk Cutaneous Amyloidosis, 924 Norman Levine & Cynthia Burk Dermatosis Papulosa Nigra, 928 Norman Levine & Cynthia Burk Ephelides (Freckles), 929 Norman Levine & Cynthia Burk Erythema ab Igne, 931 Norman Levine & Cynthia Burk viii

Erythema Dyschromicum Perstans, 933 Norman Levine & Cynthia Burk Erythromelanosis Follicularis Faciei et Colli, 935 Norman Levine & Cynthia Burk Erythrose Péribuccale Pigmentaire of Brocq, 937 Norman Levine & Cynthia Burk Extracutaneous Neuroendocrine Melanoderma, 938 Norman Levine & Cynthia Burk Felty Syndrome and Rheumatoid Arthritis, 939 Norman Levine & Cynthia Burk Hyperpigmentation Associated with Human Immunodeﬁciency Virus (HIV) Infection, 941 Philippe Bahadoran Melanoacanthoma, 946 Norman Levine & Cynthia Burk Phytophotodermatitis, 948 Norman Levine & Cynthia Burk Polyneuropathy, Organomegaly, Endocrinopathy, M Protein, and Skin Changes: POEMS Syndrome, 951 James J. Nordlund Urticaria Pigmentosum and Mastocytosis, 954 James J. Nordlund Poikiloderma of Civatte, 959 Vlada Groysman & Norman Levine Riehl’s Melanosis, 961 Scott Bangert & Norman Levine Atrophoderma of Pasini et Pierini, 963 James J. Nordlund, Norman Levine, Charles S. Fulk, & Randi Rubenzik Hyperpigmentation Associated with Scleromyxedema and Gammopathy, 965 Kazunori Urabe, Juichiro Nakayama, & Yoshiaki Hori Ichthyosis Nigricans, Keratoses, and Epidermal Hyperplasia, 965 James J. Nordlund Morphea and Scleroderma, 967 James J. Nordlund Pigmentary Changes Associated with Addison Disease, 969 Cindy L. Lamerson & James J. Nordlund Pigmentary Changes Associated with Cutaneous Lymphomas, 972 Debra L. Breneman 51 Hypermelanosis Associated with Gastrointestinal Disorders, 979 Porphyria Cutanea Tarda, 979 Eun Ji Kwon & Victoria P. Werth Cronkhite–Canada Syndrome, 983 Eun Ji Kwon, James J. Nordlund, & Victoria P. Werth Hemochromatosis and Hemosiderosis, 986 Joerg Albrecht & Victoria P. Werth Primary Biliary Cirrhosis, 992 Joerg Albrecht & Victoria P. Werth Inﬂammatory Bowel Disease and Pigmentation, 995 Joerg Albrecht & Victoria P. Werth

CONTENTS

Pellagra, 995 Eun Ji Kwon & Victoria P. Werth Peutz–Jeghers Syndrome, 999 Nancy Burton Esterly, Eulalia Baselga, & Beth A. Drolet 52 Acquired and Congenital Dermal Hypermelanosis, 1003 Sacral Spot of Infancy, 1003 Sang Ju Lee, Seung Kyung-Hann, & Sungbin Im Nevus of Ota, 1006 Sang Ju Lee, Seung Kyung-Hann, & Sungbin Im Nevus of Ito, 1012 Sang Ju Lee, Seung Kyung-Hann, & Sungbin Im Phakomatosis Pigmentovascularis, 1013 Sang Ju Lee, Seung Kyung-Hann, & Sungbin Im Other Congenital Dermal Melanocytosis, 1015 Sang Ju Lee, Seung Kyung-Hann, & Sungbin Im Acquired Dermal Melanocytosis, 1016 Sang Ju Lee, Seung Kyung-Hann, & Sungbin Im Carleton–Biggs Syndrome, 1017 Sang Ju Lee, Seung Kyung-Hann, & Sungbin Im Acquired Bilateral Nevus of Ota-like Macules (ABNOM), 1017 Sang Ju Lee, Seung Kyung-Hann, & Sungbin Im Blue Macules Associated with Progressive Systemic Sclerosis, 1018 Sang Ju Lee, Seung Kyung-Hann, & Sungbin Im 53 Mixed Epidermal and Dermal Hypermelanoses and Hyperchromias, 1020 Melasma, 1020 Sang Ju Lee, Seung Kyung-Hann, & Sungbin Im Melanosis from Melanoma, 1023 Sang Ju Lee, Seung Kyung-Hann, & Sungbin Im 54 Drug-induced or -related Pigmentation, 1026 Peter A. Lio & Arthur J. Sober Section 6: Disorders of Pigmentation of the Nails and Mucous Membranes, 1055 55 The Melanocyte System of the Nail and its Disorders, 1057 Robert Baran, Christophe Perrin, Luc Thomas, & Ralph Braun

56 Pigmentary Abnormalities and Discolorations of the Mucous Membranes, 1069 John C. Maize, Jr. & John C. Maize, Sr. Section 7: Benign Neoplasms of Melanocytes, 1091 57 Common Benign Neoplasms of Melanocytes, 1093 Pigmented Spindle Cell Nevi, 1093 Julie V. Schaffer & Jean L. Bolognia Speckled Lentiginous Nevus (Nevus Spilus), 1098 Julie V. Schaffer & Jean L. Bolognia Melanocytic (Nevocellular) Nevi and Their Biology, 1112 Julie V. Schaffer & Jean L. Bolognia 58 Rare Benign Neoplasms of Melanocytes, 1148 Nevus Aversion Phenomenon, 1148 James J. Nordlund Melanotic Neuroectodermal Tumor of Infancy, 1148 Julie V. Schaffer & Jean L. Bolognia Pilar Neurocristic Hamartoma, 1157 Julie V. Schaffer & Jean L. Bolognia Section 8: Treatment of Pigmentary Disorders, 1163 59 Topical Treatment of Pigmentary Disorders, 1165 Rebat M. Halder & James J. Nordlund 60 Chemophototherapy of Pigmentary Disorders, 1175 Rebat M. Halder & James J. Nordlund 61 UVB Therapy for Pigmentary Disorders, 1183 Thierry Passeron & Jean-Paul Ortonne 62 Sunscreens and Cosmetics, 1188 James J. Nordlund & Rebat M. Halder 63 Surgical Treatments of Pigmentary Disorders, 1191 Rebat M. Halder & James J. Nordlund 64 Laser Treatment of Pigmentary Disorders, 1198 Rebat M. Halder, Lori M. Hobbs, & James J. Nordlund Index, 1205 Color Plates, 1230

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Contributors

Zalfa Abdel-Malek

PhD Research Professor of Dermatology, Department of Dermatology, University of Cincinnati College of Medicine, Cincinnati, OH, USA

Joerg Albrecht MD Department of Dermatology, University of Pennsylvania, Philadelphia, PA, USA Mayra Alvarez-Franco

(deceased) Formerly of the Departments of Dermatology and Pathology, Yale University School of Medicine, New Haven, CT, USA

Joseph T. Bagnara PhD Professor Emeritus, Department Cell Biology and Anatomy, University of Arizona College of Medicine, Tucson, AZ, USA Philippe Bahadoran

MD PhD Assistant Professor of Medicine, Service de Dermatologie, Hôpital l’Archet, Nice, France

Debdutta Bandyopadhyay

PhD Instructor of Dermatology, Hufﬁngton Center on Aging, Baylor College of Medicine, Houston, TX, USA

Raymond E. Boissy PhD Professor of Dermatology and Cell Biology, Department of Dermatology, University of Cincinnati College of Medicine, Cincinnati, OH, USA Jean L. Bolognia MD Professor of Dermatology, Department of Dermatology, Yale University School of Medicine, New Haven, CT, USA Barbara Boone MD Resident in Dermatology, Universitair Ziekenhuis Ghent, Ghent, Belgium Ralph P. Braun

MD Assistant Professor, Pigmented Skin Lesion Unit, Department of Dermatology, University Hospital, Geneva, Switzerland

Debra L. Breneman

MD Associate Professor of Dermatology, University of Cincinnati, Cincinnati, OH, USA

Stefania Briganti

PhD San Gallicano Dermatological Institute, Rome, Italy

Scott D. Bangert

MD Banner Good Samaritan Hospital, Department of Medical Education, Phoenix, AZ, USA

Murray H. Brilliant PhD Lindholm Professor of Genetics, Department of Pediatrics, University of Arizona College of Medicine, Tucson, AZ, USA

Robert Baran MD Head of Nail Disease Center, Cannes, France

Lieve Brochez

Gregory S. Barsh MD PhD Professor of Genetics and Pediatrics, Beckman Center for Molecular and Genetic Medicine, Stanford University School of Medicine, Stanford, CA, USA

Cynthia J. Burk

Eulalia Baselga

Stella D. Calobrisi

MD Consultant in Pediatric Dermatology, Hospital de la Santa Creu I Sant Pau, Barcelona, Spain

Dorothy C. Bennett

MA PhD Professor of Cell Biology, St George’s Hospital, University of London, London, UK

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MD PhD Professor of Dermatology, Department of Dermatology, University Hospital of Ghent, Ghent, Belgium

MD Section of Dermatology, University of Arizona Health Sciences Center, Tucson, AZ, USA MD Medical Director, The Dermatology Clinic, Boca Raton, FL, USA

Tania Cestari

MD PhD Associate Professor of Dermatology, Department of Dermatology, University of Rio Grande do Sul, Hospital de Clínicas de Porto Alegre, Brazil

CONTRIBUTORS

Henry H. L. Chan MBBS MSc (Clin Derm) MD FRCP FRCP(Ed) FRCP(Glasg) FHKCP FHKAM (Med) Specialist in Dermatology and Honorary Clinical Associate Professor, University of Hong Kong and Chinese University of Hong Kong, Hong Kong Peter Man Hon Chan MD Clinical Research Fellow, University of Minnesota, Eagan, MN, USA Peggy L. Chern MD Department of Dermatology, University of Iowa Hospitals and Clinics, Iowa City, IA, USA Mary K. Cullen

MD Department of Cell Biology and Physiology, Clinical Instructor, Washington University School of Medicine, St Louis, MO, USA

José C. García-Borrón PhD Professor of Biochemistry and Molecular Biology, School of Medicine, University of Murcia, Murcia, Spain Rosemary J. Geary MD President, East Valley Dermatology Center, Chandler, AZ, USA Vanessa C. Gray-Schopfer PhD Signal Transduction Team, Cellular and Molecular Biology Section, Institute of Cancer Research, London, UK Pearl E. Grimes MD Vitiligo and Pigmentation Institute of Southern California, David Geffen School of Medicine, University of California at Los Angeles, Los Angeles, CA, USA Vlada Groysman

Beth A. Drolet

MD Section of Dermatology, University of Arizona Health Sciences Center, Tucson, AZ, USA

Carol A. Erickson

Ruth Halaban PhD Senior Research Scientist, Department of Dermatology, Yale University School of Medicine, New Haven, CT, USA

MD Department of Dermatology, The Medical College of Wisconsin, Milwaukee, WI, USA PhD Professor of Molecular Cellular Biology, University of California at Davis, Davis, CA, USA

Nancy B. Esterly MD Professor of Dermatology and Pediatrics, Department of Dermatology, The Medical College of Wisconsin, Milwaukee, WI, USA Dong Fang MD PhD Staff Scientist, The Wistar Institute, Philadelphia, PA, USA Sharon A. Foley MD Department of Dermatology, Washington University School of Medicine, St Louis, MO, USA Peter S. Friedmann MD FRCP FMedSci Professor of Dermatology and Head of Division, Dermatopharmacology Unit, Southampton General Hospital, Southampton, UK Charles S. Fulk

MD FACP Department of Pathology, Vanderbilt University, Nashville, TN, USA

Rebat M. Halder

MD Professor and Chairman, Department of Dermatology, Howard University, Washington, DC, USA

Seung-Kyung Hann MD PhD Director, Korea Institute of Vitiligo Research, Yongsan-Gu, Seoul, Korea Rudolf Happle MD Professor of Dermatology, Department of Dermatology, Philipp’s University of Marburg, Marburg, Germany Vincent J. Hearing

PhD Chief, Pigment Cell Biology Section, Laboratory of Cell Biology, National Institutes of Health, Bethesda, MD, USA

Meenhard Herlyn DVM, DSc Professor and Chair of Program of Molecular and Cellular Oncogenesis, The Wistar Institute, Philadelphia, PA, USA Stan P. Hill

MD Denver, CO, USA

Anne-Sophie J. Gadenne MD Department of Dermatology, University of Cincinnati College of Medicine, Cincinnati, OH, USA

Lori M. Hobbs MD

William A. Gahl MD PhD Clinical Director, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, USA

Thomas J. Holstein

Sheila S. Galbraith MD Assistant Professor of Dermatology, Dermatology Department, Medical College of Wisconsin, Milwaukee, WI, USA

Assistant Professor, Department of Dermatology, Howard University, Washington, DC, USA PhD Professor Emeritus, Roger Williams University, Bristol, RI, USA

Yoshiaki Hori

MD PhD (deceased) Formerly Honorary Professor, Department of Dermatology, Kyushu University, Fukuoka, Japan

xi

CONTRIBUTORS

Thomas J. Hornyak MD PhD Dermatology Branch, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA

I. Caroline Le Poole

Marjan Huizing

Norman Levine MD Professor of Medicine, Section of Dermatology, University of Arizona College of Medicine, Tucson, AZ, USA

PhD Head, Cell Biology of Metabolic Disorders Unit, Medical Genetics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, USA

PhD Loyola University Medical Center Department of Pathology, Cancer Center, Maywood, IL, USA

Peter A. Lio Sungbin Im MD PhD Director, Kangnam Wootaeha Skin and Esthetic Clinic, Yeoksam-Dong, Kangnam-Gu, Seoul, Korea Genji Imokawa

PhD Director, Skin-Bio Assess Institute, Utsunomiya, Tochigi, Japan

Shosuke Ito

PhD Professor of Chemistry, Department of Chemistry, Fujita Health University School of Health Sciences, Toyoake, Aichi, Japan

Ana Luisa Kadekaro

PhD Research Instructor of Dermatology, University of Cincinnati College of Medicine, Cincinnati, OH, USA

Richard A. King MD PhD Director, Genetics Division, Department of Medicine and Associate Director, Institute of Human Genetics, University of Minnesota, Minneapolis, MN, USA Sidney N. Klaus

MD Professor of Dermatology, Dartmouth-Hitchcock Medical Center, Lebanon, NH, USA

Wolfgang Küster MD Professor and Chief of Dermatology, TOMESA Clinic, Department of Dermatology, Bad Salzschlirf, Germany Eun Ji Kwon

BSc Department of Dermatology, University of Pennsylvania, Philadelphia, PA, USA

Jean-Philippe Lacour

MD Service de Dermatologie, Hôpital Archet 2, Nice, France

Cindy L. Lamerson MD Clinical Assistant Professor, University of Nevada, Reno, NV, USA

MD Instructor in Dermatology, Beth Israel Deaconess Medical Center, Boston, MA, USA

John C. Maize Sr MD Medical Director, DermPath Diagnostics, Maize Center for Dermatopathology, Mt Pleasant, SC, USA John C. Maize Jr

MD DermPath Diagnostics, Maize Center for Dermatopathology, Mt Pleasant, Clinical Assistant Professor of Dermatology, Medical University of South Carolina, SC, USA

Susan Bayliss Mallory MD FAAD FAAP Professor of Internal Medicine (Dermatology) and Pediatrics, Division of Dermatology, Washington University School of Medicine, St Louis, MO, USA Jiro Matsumoto

PhD Emeritus Professor, Department of Biology, Keio University, Kohoku-ku, Yokohama, Japan

Estela E. Medrano PhD Professor of Molecular and Cellular Biology and Dermatology, Hufﬁngton Center on Aging, Baylor College of Medicine, Houston, TX, USA Henk E. Menke MD PhD Dermatologist, Department of Dermatology, Sint Franciscus Gasthuis, Rotterdam, The Netherlands Allan D. Mineroff MD Lansdale, PA, USA

Gisela E. Moellmann

PhD Professor Emeritus, Yale University School of Medicine, New Haven, CT, USA

Jean-Marie Naeyaert

MD PhD Professor of Dermatology and Head, Department of Dermatology, University Hospital of Ghent, Ghent, Belgium

Edward J. Land

BSc PhD Honorary Professor, Lennard-Jones Laboratories, School of Geographical and Physical Sciences, Keele University, Keele, UK

Juichiro Nakayama

Sang Ju Lee MD PhD Director, Yonsei Star Skin and Laser Clinic, Chancheon-Dong, Seodaemun-Gu, Seoul, Korea

Marcelius Davy Njoo

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MD PhD Department of Dermatology, School of Medicine, Fukuoka University, Fukuoka, Japan

MD PhD Streekzierenhuis Middentwente, Hengelo, The Netherlands

CONTRIBUTORS

James J. Nordlund MD Dermatologist and Professor Emeritus, Group Health Associates, Cincinnati, OH, USA

Patrick A. Riley

William S. Oetting PhD Associate Professor, Genetics Division, Department of Medicine, Institute of Human Genetics, University of Minnesota, Minneapolis, MN, USA

Randi E. Rubenzik

Jean-Paul Ortonne

MD Professor of Dermatology and Chairman, Department of Dermatology, University of Nice-Sophia Antipolis, Nice, France

Monica Ottaviani PhD San Gallicano Dermatological Institute, Rome, Italy

MBBS PhD DSc FRCPath CBiol FIBiol FLS Professor Emeritus of Cell Biology and Director, Totteridge Institute for Advanced Studies, London, UK MD FAAD

Sun City West, AZ, USA

Tadeusz Sarna

PhD DSc Professor of Biophysics, Department of Biophysics, Faculty of Biotechnology, Jagiellonian University, Krakow, Poland

Julie V. Schaffer MD Assistant Professor of Dermatology and Pediatrics, New York University School of Medicine, New York, NY Glynis A. Scott

David M. Parichy

MD Associate Professor of Dermatology and Pathology, Department of Dermatology, University of Rochester School of Medicine, Rochester, NY, USA

Thierry Passeron

MD Clinical Assistant in Medicine, Nice, France

Vijayasaradhi Setaluri PhD Associate Professor of Dermatology, University of Wisconsin School of Medicine and Public Health, Madison, WI, USA

Christophe Perrin

MD Laboratoire d’Anatomie Pathologique, Hôpital L. Pasteur, Nice, France

Anita P. Sheth

Mauro Picardo

Pranav B. Sheth MD Assistant Professor, Department of Dermatology, University of Cincinnati, Cincinnati, OH, USA

PhD Assistant Professor, Department of Biology, University of Washington, Seattle, WA, USA

MD San Gallicano Dermatological Institute, Rome, Italy

Tanusin Ploysangam

MD PhD Director, Institute of Beauty and Health Science, and Director, Belage Skin Care and Cosmetic Center, Nunthawan Prachacheun, Bangtalad, Nonthaburi, Thailand

Walter C. Quevedo Jr PhD Emeritus Professor of Biology, Department of Molecular and Cell Biology and Biochemistry, Brown University, Providence, RI, USA Christopher A. Ramsden BSc PhD DSc Cchem FRSC Professor of Organic Chemistry, Lennard-Jones Laboratories, School of Geographical and Physical Sciences, Keele University, Keele, UK

H. Randolph Byers MD PhD Professor of Dermatology, Boston University School of Medicine, Boston, MA, USA Mark V. Reedy

PhD Assistant Professor of Biology, Creighton University, Omaha, NE, USA

Germaine N. Relyveld

MD Department of Dermatology, Academic Medical Center, Amsterdam, The Netherlands

MD Department of Dermatology, University of Cincinnati College of Medicine, Cincinnati, OH, USA

Shigeki Shibahara MD PhD Professor, Department of Molecular Biology and Applied Physiology, Tohoku University School of Medicine, Aoba-ku, Sendai, Miyagi, Japan Arthur J. Sober MD Professor of Dermatology, Harvard Medical School, Massachusetts General Hospital, Boston, MA, USA Francisco Solano PhD Professor of Biochemistry and Molecular Biology, School of Medicine, University of Murcia, Spain Richard A. Spritz, MD Professor and Director, Human Medical Genetics Program, University of Colorado Health Sciences Center at Fitzsimons, Aurora, CO, USA Harold M. Swartz MD PhD Professor of Radiology, Dartmouth Medical School, Hanover, NH, USA Kazuhisa Takeda

MD PhD Assistant Professor of Molecular Biology, Department of Molecular Biology and Applied Physiology, Tohoku University School of Medicine, Aoba-ku, Sendai, Miyagi, Japan

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CONTRIBUTORS

Luc Thomas MD PhD Professor and Chairman, Department of Dermatology, Lyon 1 University, Hotel Dieu, Lyon, France

Victoria P. Werth MD Professor of Dermatology and Chief, VA Dermatology, Department of Dermatology, University of Pennsylvania, Philadelphia, PA, USA

Marnie D. Titsch MD Dermatology Specialist, Oyster Point Dermatology Inc., Newport News, VA, USA

Wiete Westerhof

Peggy Tong MD Dermatology Associates, Milwaukee, WI, USA Kazunori Urabe

MD PhD Associate Professor, Department of Dermatology, Graduate School of Medical Science, Kyushu University, Fukuoka, Japan

Kazumasa Wakamatsu

PhD Professor, Fujita Health University School of Health Sciences, Toyoake, Aichi, Japan

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MD PhD Assistant Professor of Dermatology, Netherlands Institute for Pigment Disorders and Department of Dermatology, Academic Medical Center, University of Amsterdam, Amsterdam, The Netherlands

Antony R. Young PhD Professor of Experimental Photobiology, King’s College London and St John’s Institute of Dermatology, St Thomas’ Hospital, London, UK

Foreword

In this second edition of The Pigmentary System: Physiology and Pathophysiology, the editors have expanded upon the dynamic ﬁeld of pigment cell biology to reﬂect not only the rapidly moving pace of developments in this broad ﬁeld but also the explosive growth of research and development across the basic life sciences. We have all beneﬁted enormously from the recent growth in the research budgets of agencies such as the National Institutes of Health (NIH), and these beneﬁts transcend beyond grant recipients to citizens who beneﬁt from better treatments or management protocols for diseases that in the past were nearly always seriously debilitating. Never before have advances come so rapidly or with so much promise for improving the human condition. These improvements come as a result of better understanding of all aspects of the composition and function of living cells. From the molecules that direct activity to those that confer shape and phenotype, we know more today about the basic building blocks of living matter than at any time previously in our history. Perhaps even more astounding is the pace at which discoveries in the life sciences continue to be made. In recent years we have experienced near-exponential growth in the quantity and quality of information available across all sectors of the life sciences. There is little doubt that as we move toward the end of the ﬁrst decade of the twenty-ﬁrst century we will have answers to many questions, some of which we never imagined being able to ask. The pigmentary system, as a model system for studying basic questions about differentiating stem cells, has provided ample opportunities for study across a broad range of inter-

ests and expertise. Animal pigment has long been known to function protectively in some animals as well as decoratively in others. Pigment can provide camouﬂage for some and ﬂamboyant advertisement for others. The embryonic origin of skin pigment cells, the neural crest, suggests that pigment cells share common traits with a wide variety of nerve cells. And yet, like all stem cells, neural crest cells can differentiate along a multitude of pathways, perhaps to become a sensory neuron or to become a melanocyte. These highly motile embryonic cells, even after settling down and differentiating, still retain a remarkable capacity for further motion and differentiation or reversion. Witness the havoc that can ensue if melanocytes become cancerous and migrate to all parts of the body to form tumors. The ﬁrst edition of The Pigmentary System was unquestionably the deﬁnitive text of its time. The editors, who are themselves pre-eminent in this ﬁeld, have worked with the authors to provide signiﬁcant revisions to nearly every chapter, and a new chapter on melanocytes as malignant cells has been added. Virtually any topic related to pigmentary systems can be found in this single source, from animal chromatophores to human pigmentary disorders, from basic cell biology to clinical studies and results. Congratulations to the editors and to those researchers who are actively engaged in helping us better understand the complexities of this fascinating system! Sally Frost Mason PhD Purdue University West Lafayette, IN, USA

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Preface

This second edition of The Pigmentary System: Physiology and Pathophysiology presents the latest information about the biology of melanocytes and their disorders. The ﬁrst section is dedicated to the science of pigmentation and has been completely rewritten, not just updated. The chapters have been reorganized to provide the most current and comprehensive review of the science of pigmentation. New chapters have been added and some old chapters in the ﬁrst edition have been deleted. All of these were chapters on methodology that were rapidly outdated after publication. Some information and data remain constant and are unchanged in this edition. Other topics are entirely new or signiﬁcantly enlarged since publication of the previous edition. Part II on the pathophysiology of pigmentary disorders is mostly updated. New chapters have been included. A lexicon on proper terminology for description of skin color has been included as the ﬁrst chapter in this section. Unless the scientiﬁc and clinical communities use the same terminology to describe and identify scientiﬁc or clinical topics, there will be confusion about the nature and treatment of pigmentary or dyschromic disorders. We hope that this lexicon becomes standard in the dermatological and scientiﬁc communities. The comments that we made in the preface to the ﬁrst edition are still very relevant. A few are worth repeating. This book is intended to be a reference book for both the scientiﬁc and the clinical communities. The fact that there is so much

ongoing scientiﬁc work related to pigmentation is an indicator of the importance of melanocytes. It also is a clue that melanocytes, melanotropins and related cells and factors are not just a mechanism for sun protection. It is obvious and certain that melanocytes and melanin are protective against sunlight. But the role of melanin in the eyes and ears is more than just sun protection. Melanocytes are involved in many processes within the epidermis. Melanotropins have effects on virtually every organ system in the body and control inﬂammation, appetite, some functions within the nervous system, and other critical functions. The pigmentary system, including melanin, melanocytes and melanotropins, is best described as a modulator that helps the body adapt to the environmental changes related to seasonal ﬂuctuations in sunlight, temperature, and humidity. We hope that scientists, clinicians, students, and residents ﬁnd this book useful in their studies on skin color, melanocytes, and melanin, and that it expands their perspective and view about the pigmentary system as a modulator, not just a method of sun protection. James J. Nordlund Raymond E. Boissy Vincent J. Hearing Richard A. King William S. Oetting Jean-Paul Ortonne

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Acknowledgments

The editors wish to acknowledge the very gracious and generous support of the following companies. Without their support, this project would not have been possible. Galderma International, Paris, France Proctor and Gamble, Cincinnati, OH, USA Combe Inc., White Plains, NY, USA

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The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

I

The Physiology of the Pigmentary System

The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

1

Historical and Comparative Perspectives of the Pigmentary System

The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

1

A History of the Science of Pigmentation Sidney N. Klaus

Introduction In Europe, prior to the seventeenth century, notions about the origin of human skin color were based largely on myths and fanciful stories passed down from the ancient world. The stories focused on providing explanations for the blackness of Africans. Europeans with an ethnocentric perspective thought that it was necessary to justify the darkness of outsiders rather than to explain their own paleness. In Europe, two theories about the origin of Africans’ color predominated. The ﬁrst proposed that the black hue of African skin was a consequence of the intense sunlight and heat to which Africans were exposed. The second idea proposed that African color was a result of divine intervention, a concept that stemmed from the well-known passage in Genesis describing Noah’s curse on Ham and his progeny. Such traditional views of skin color were questioned during the Age of Discovery. Explorers were returning home from remote regions of the globe and bringing with them descriptions of black-, yellow-, or red-hued peoples. The array of colors fascinated scientists and excited them to investigate how these skin colors originated. During the sixteenth century, the scientiﬁc community prepared for this task. Scientists learned new techniques of human dissection and new methods of chemical analysis. More importantly, they adopted a new philosophical approach for the study of natural phenomena. Rather than analysis of phenomenon based on a priori assumptions, they based their analyses on ﬁrsthand observations and experimentation. By the seventeenth century, anatomists and physicians had begun to collect objective data about skin color. For the next two centuries, so many important observations had been collected about the seat of skin color and the causes of its diversity that, as one researcher boasted, “a good sized volume was scarcely large enough to contain them.” The furious pace of forward progress was slowed somewhat in the early part of the nineteenth century when skin color scientists in both Europe and America were drawn into acrimonious debates over social issues, especially slavery and the place of “peoples of color” in the family of man. By the end of the century, the slavery issue had been settled, and the discovery of the cell and improved methods for the microscopic examination of the skin had stimulated the discipline to set off in new directions.

In retrospect, it is clear that the science of skin color had completed its early phase of development by the 1840s. With the discovery of the cellular nature of the epidermis, the “modern” phase of skin color science had begun (Meirowsky, 1940). This chapter brieﬂy summarizes the ﬁrst 250 years of the earlier phases of pigment research, from its birth in the 1600s to its “coming of age” in the 1840s. A history of the modern era, from the 1840s to the present, may be found in reviews by Becker (1959) and Nordlund et al. (1989).

Early Anatomic Discoveries To the ancient Greeks and Romans who were acquainted with the peoples of Ethiopia, the black color of the Africans was regarded as their most characteristic and most curious feature. Greek and Roman scientists were not equipped to investigate the causes of dark complexions in a meaningful way. They understood little about dissection of individual tissues and were unable to sort out the anatomic details related to the skin. They regarded the skin as an amorphous membrane made from the congealing of “moist exhalations” driven to the surface of the body by internal heat. They believed that the color of skin was imposed either from outside by the sun or by some inner humor. These notions about skin color persisted through the Middle Ages and into the Renaissance. In 1543, Vesalius succeeded in splitting the skin into two layers by applying a burning candle to the abdomen of a living subject. He did not examine the skin layers for their content of color. It was not until 75 years later that the ﬁrst “scientiﬁc” study of skin color was performed. In 1618, Jean Riolan the younger, a Parisian anatomist, was the ﬁrst person to make a detailed analysis of skin color. He used a technique similar to that of Vesalius to separate the skin of a black subject into two layers. But he used a vesicant rather than a ﬂame to produce a blister and carefully examined the two layers for their color and texture. He found that, while the top of the blister (the cuticle) was pigmented, the base (the cutis) remained white. “Color . . .” he announced “. . . lay in the outer layer, and did not go so deep as the true skin” (Riolan, 1618). To Riolan, this ﬁnding reinforced the idea that heat and sunlight were the ultimate causes of blackness. Riolan reasoned that, if the sun caused skin darkening, it was expected that the outer cuticle would be darker than the inner cutis. 5

CHAPTER 1

A few years later, Alexander Read, an anatomist working in London, repeated Riolan’s experiments and found the same results. However, Read offered a different interpretation. The skin pigment, he concluded, was not caused by singeing the body’s surface. He concluded rather that skin color was derived from the body’s inner humors. As they escaped through the skin, some of the elements became attached to the surface and dried, thus producing the pigment. Negroes, he argued, had darker skin because they had larger pores than whites, a morphologic variation allowing greater volumes of their humors to escape leaving behind larger amounts of black “remnants” of the humors (Read, 1642). Perhaps the most detailed analysis of skin color in the seventeenth century was carried out by the distinguished writer and physician, Sir Thomas Browne. Browne had listened carefully to the accounts of explorers who had found variously colored races, and he concluded from these accounts that color did not arise from the inﬂuence of climate. “If the fervor of the Sun were the cause of the Negroes’ complexion, . . .” he wrote, “. . . it is reasonable to assume that inhabitants of the same latitude, subjected unto the same vicinity of the Sun, partake of the same hue and complexion, which they do not.” In addition, he noted that Africans who had been transplanted into “. . . cold and phlegmatic habitations . . .” had not turned white, but “. . . continued their hue, both in themselves and their generations.” He rejected the idea that skin blackness was acquired from a “. . . power of imagination . . .” or from “. . . anointing with bacon and fat substances.” Instead, he thought that the tincture of the skin was based on an inborn trait, a trait that was passed from father to son by the sperm. Browne was not an armchair scientist. His diary records the outline of numerous experiments he carried out to determine for himself the seat of color. For example, he set out to make a “. . . blistering plaster in a negroes skinne and trie if the next skin will bee white . . .” and to see whether a “. . . vesication will do anything upon a dead cold body.” He even urged his son Edward, a medical student, to carry out additional studies. He wrote to him asking that he “. . . separate the skin of a black person with boiling water and take notice of the cuticula and cutis and observe how the scarres become whiter or less blacke than any other parts.” He also urged him to “. . . touche the skin with aqua fortis and see how it will alter the colour.” Unfortunately, Browne did not record the results of these experiments (Keynes, 1964). Another seventeenth century polymath, Robert Boyle, devoted a chapter of his book Experiments and Considerations Touching Colours to what he called “the Cause of the Blackness of those many nations which by one common Name we are wont to call Negroes.” He examined the various causes proposed at that time and denied that either Noah’s curse on Ham or “heat alone” could produce a true blackness. Instead, he focused on principles of inheritance, concluding that the “Principal Cause of the Blackness of Negroes is some Peculiar and Seminal Impression.” As evidence, he reported that “the off-spring of Negroes, trans6

planted out of Africa, retain still the complexion of their Progenitors” (Boyle, 1664).

Malpighi’s Innovation: the Rete Mucosum Riolan’s contention that the cuticle was the seat of color had been in place for less than a century when it was superseded by an entirely different notion proposed by the Italian anatomist, Marcello Malpighi. In 1667, Malpighi stated that the seat of color was not the cuticle. Rather, he proclaimed that color was located in a separate layer of the skin sandwiched between the cuticle and the cutis. He named this layer the rete mucosum. The discovery of rete came about in the following way. In 1665, Malpighi had been searching for the structures that mediated the sensation of taste, and he succeeded in ﬁnding large papillae just below the surface of the tongue. Turning to the skin to see if he could ﬁnd similar structures that mediated touch, he found that he was unable to separate the skin’s layers properly by boiling, the method that he had used previously. He was forced to use putrefaction, an old dissection technique in which the separation of parts is enhanced by allowing a small piece of tissue to decompose partially. Using this method, he prepared skin from cadavers and found that he could easily lift the cuticle from the cutis. By this method, he was able to identify the dermal papillae. However, he also found a mucoid material draped over the papillae, which he regarded as a protective covering. The material had a netlike appearance when seen from above, so he called it the rete mucosum. Malpighi’s experiments were not directed at discovering the proximate cause of skin color. However, in one experiment in which he used skin from an Ethiopian cadaver, he noted that the mucoid material was tinged black. On the basis of this ﬁnding, he later wrote “It is certain that the cutis (of the Ethiopian) is white, as is the cuticula too; hence all their blackness arises from the underlying mucous and netlike body” (Malpighi, 1665). Malpighi did not follow up on his discovery of the “seat of color” but the putrefaction technique he used allowed the layers of the skin to be separated gently. The method produced reliable results, and other investigators soon expanded on Malpighi’s ﬁnding. In 1677, three years after Malpighi published his ﬁndings, Johann Pechlin, a Dutch anatomist, used Malpighi’s method of dissection to make a thorough study of Ethiopian skin. In one experiment, after he lifted off the cuticle, he scraped away the black mucous and reapplied the scarf to the cutis. He reported that “. . . a whiteness came forth immediately of the sort seen often enough in Europe. In truth the surface of the skin was scarcely unattractive.” Pechlin was concerned that his results obtained using cadaver skin might not be the same had he studied living skin. To his dismay, he was unable to convince living subjects to cooperate in his experiments “. . . because of I don’t know what sort of evil on the part of the Negroes” (Pechlin, 1677).

A HISTORY OF THE SCIENCE OF PIGMENTATION

A few years later, the Dutch microscopist Antoni van Leeuwenhoek, in a letter to the Royal Society of London, reported on his observations made using his microscope of the skin of a black Moorish girl. “I took from several parts of the arms the outer skin with a ﬁne little instrument and found that it consisted of little scales. Putting these scales before my microscope I found them to be not as transparent as those of my skin.” Van Leeuwenhoek concluded that the black color of the skin was the result of the black scales, and that “the little vessels which form the scales of the Moors” may possibly develop a slightly darker color (Collected Letters of Antoni van Leeuwenhoek, 1683–1684; see Schierbeek, 1952). If one follows Malpighi’s directions for uncovering the rete, as the cuticle is lifted off one observes tiny mucoid threads bridging the angle between the two layers. William Hunter described them as “. . . an inﬁnite number of ﬁlaments, as ﬁne as the most delicate threads of a spider’s web, that pass between the cutis and the more external integuments” (Hunter, 1764). According to some anatomists, the threads were tiny vessels. This idea led some of the more inventive investigators to propose that the coloring matter in Africans’ skin was not the mucous material itself but was a black ﬂuid contained within the web of small vessels. One contemporary observer wrote “. . . between the outward and inner skin of the corpse can be found a kind of vascular plexus, spread over the whole body like a web or net, which was ﬁll’d with a Juice as black as Ink” (Marana, 1801). It became something of a challenge to prove unequivocally that these ﬁlaments were in fact “vessels.” In anatomic laboratories, the identiﬁcation of small vessels in anatomic preparations relied on their visualization after they had been injected with colored glue or isinglass. Only a few investigators were skilled enough to inject these “. . . most delicate threads.” One anatomist who was given credit for the ﬁrst demonstration of this vascular network was William Baynham, an American who had moved to London in 1769. Although Baynham never published his ﬁndings, he placed his specimens on exhibition in John Hunter’s anatomic museum. Not all Baynham’s colleagues were convinced by his discovery. William Cruickshank made a careful study of Baynham’s preparations and later wrote that he was “. . . not perfectly satisﬁed” with them. He concluded that Baynham had not injected the vessels of the rete, but rather a series of vessels that lay between the rete and the cutis (Cruickshank, 1795). In the eighteenth century, the attention of researchers began to move from the anatomy of the rete to the “nature” of the coloring matter itself. Early attempts to collect and analyze material from the rete were not productive. Alexis Littre, a French surgeon, soaked pieces of skin from a Negro cadaver for a week either in warm water or in spirits of wine but was unable to extract any of the coloring matter (Littre, 1720). A few years later, another French scientist, Pierre Barrere, in an essay submitted to the Academy of Bordeaux in 1741, denied Malpighi’s claim that the pigment of Africans’ skin arose from the “corps reticulaire.” It is evident, he wrote, the coloring matter in the skin of Africans was bile. He reported

that he had examined cadavers of Africans and observed that “the bile is black, and the blackness of the skin is in proportion to the blackness of the bile.” Later in the century, another investigator attempted to prove that the pigment of Africans was derived from bile. Samuel Stanhope Smith, a professor of moral theology at the College of New Jersey, had learned from an American colleague that bile exposed to the sun and air changed its color to black. Smith hypothesized that, in southern climates, the secretion of bile was augmented and, when the bile reached the skin, it became “. . . more languid and almost ﬁxed.” In the skin, the aqueous parts of the bile easily escape through the pores of the skin by perspiration, while the more dense portion remained in a glutinous state within the rete. There, it received the repeated radiation from the sun and atmosphere and turned black. Smith believed that a cold environment would reverse the process of darkening and render the complexion “. . . clear and ﬂorid.” According to Smith, if Africans moved to temperate climates from Africa “. . . they would soon lose their imposed color and become the color of Europeans” (Smith, 1788). Johann Friedrich Blumenbach, the German ethnologist, also proposed that bile was the coloring agent of the skin. He had learned that bile was made of a mixture of carbon and hydrogen. He suggested that, as the bile reached the skin, the hydrogen it contained became more volatile than the carbon and combined with atmospheric oxygen. This reaction left behind a black carbon residue that became embedded in the Malpighian mucous and caused the skin to darken (Blumenbach, 1865). Another “chemical” theory was advanced by the young Humphrey Davy in 1799. Davy thought that light had a great afﬁnity for oxygen and could extract it from other compounds. Davy had evidence that the mucous material of the rete was made of a colorless mixture of carbon, nitrogen, and oxygen. He proposed that sunlight extracted the oxygen from this mixture and left behind a black material made of carbon and nitrogen. To Davy, this explained differences in skin hues found throughout the world. “In northern latitudes, where the inhabitants are less exposed to light, the rete continues to contain its full proportion of oxygen, and the inhabitants are lighter in color. In the torrid zone, on the other hand, where the sun was especially intense, larger quantities of oxygen would be removed, and the blackness peculiar to the negroes will be found.” Davy had no direct proof that light could subtract oxygen from the skin, but he did cite other chemical experiments to support his claim. He reported that, when a compound that binds oxygen, such as sulfur of potash, is applied to the skin of a white person, the skin blackens. When a compound that gives off oxygen, such as muriatic acid, is applied to the skin of a Negro, it lightens (Davy, 1799). An even more imaginative notion about the nature of skin color was suggested by Immanuel Kant. Kant based his theory on the well-known capacity of phlogiston to turn blood a black color. Phlogiston was an imaginary element considered to be the essential principle of combustion. It was said to be 7

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released from the body through the lungs as part of the metabolic process. Kant believed that, in regions of the globe where the atmosphere was heavily phlogisticated, such as the coast of Africa, it would be impossible to eliminate all the body’s phlogiston by breathing. Kant proposed that the excess phlogiston would travel through the circulation and, on reaching the skin, it would precipitate in the ends of the small cutaneous vessels and turn the skin black (Lovejoy, 1959). Claude Le Cat, a French physician, claimed that the source of Africans’ blackness was neither bile nor blood. He said that the black mucous, which he called ethiops, was found only in Africans and that it was secreted directly into the skin through the tips of the cutaneous nerves (Le Cat, 1765). Although most early scientists believed that skin pigment was derived from some “internal” source such as bile or blood, a few investigators decided that the coloring matter was produced by glands within the skin itself. The earliest version of this hypothesis was offered by an English anatomist, Edward Tyson. He proposed that the black colour in Negroes’ skins came from glands that were full of a “black liquor.” He suggested that the climate might alter the glands so that “. . . they might separate from the mass of blood a differing humor from White, and by this means give a different hue to the inhabitants” (Montagu, 1943). Two French researchers, G. H. Breschet and Roussel de Vauzeme, reported that they had found a “. . . chromatogenous apparatus . . .” in the cutis which secreted the black mucous. They said that the mucous was deposited on the surface of the dermal papillae through short, excretory ducts emanating from glandular parenchyma located in the cutis (Plumbe, 1837). Another French investigator, M. Gaultier, also argued that the coloring matter was secreted within the skin but not from glands. The pigment of the skin, he said, was produced instead from hair bulbs. Gaultier’s claim was based on the results of an experiment he carried out on a living subject. He burned the skin of a Negro and then closely observed the pattern of repigmentation during the healing process. According to Gaultier, the pigment appeared ﬁrst around the “pores” through which the hairs exited and only later did it radiate out to cover the entire area of the burn (Prichard, 1813).

Differences Between Blacks and Whites As a more complete picture of skin color began to emerge by the end of the eighteenth century, political and social forces required the disciplines of anatomy and biology to take on a social dimension. Advocates of black slavery argued that Africans were not the “brothers” of Europeans but rather were the product of a separate creation. Skin color, the most obvious marker of racial identity, was forced to the center of the debate over the unity of mankind. The polygenists began looking for “signiﬁcant” anatomic differences between blacks and whites to prove they were separate species. Monogenists began looking for evidence of similarities. 8

Bolstering the polygenist cause was the English anatomist, John Gordon. Gordon claimed that whites, unlike blacks, had no rete mucosum. “I have satisﬁed myself by many dissection that in the Negro there is a Black membrane interposed between the epidermis and true skin upon which their dark color entirely depends . . . But after the strictest examination I have not been able to ﬁnd any light colored rete mucosum in the inhabitants of Great Britain, nor in those of other nations resembling them in color” (Gordon, 1815). Several anatomists disagreed. Richard Harlan, a young lecturer in anatomy from Philadelphia, wrote that “. . . the existence of the rete mucosum in the white race, so frequently denied, has been demonstrated occasionally in the European by skilful anatomists and if not deceived I myself have discovered it several times in a living European subject, by raising the epidermis with a blister, especially upon the back of the hands and the neck” (Harlan, 1835). Charles Caldwell, who had moved from Philadelphia to become a professor in a medical college in Lexington, Kentucky, reported that he too found the rete in both races, although the structures were not identical in appearance. “The rete mucosum in blacks is comparatively thick, while in the Caucasian, the rete is present but it is much thinner.” In spite of this similarity, however, Caldwell concluded that Negroes and Caucasians were sufﬁciently distinct to be called separate biological species (Caldwell, 1830).

Experiments of Nature In addition to the work carried out by anatomists and chemists in dissecting rooms and laboratories, many physicians made important contributions to the fund of information about skin color through reporting and analyzing patients with clinical problems. John Josselyn, a physician from England who was visiting the Massachusetts Bay Colony in 1675, was one of the ﬁrst physicians to advance the science of skin color by using observations based on a clinical case. While in Boston, Josselyn had been called upon to lance a “corruption” in the palm of a “Moor.” Later, he described his ﬁndings. “After I lanced it, I perceived that the Moor had one skin more than Englishmen, deeper in colour than our European veins, and upon it rests the epidermis” (Josselyn, 1675). Samuel Marcy, a physician from rural New Jersey, reported the case of an albino girl. After providing a full account of her family history, including the fact that she had two albino sisters, he went on to consider the cause of the disorder. “The mother accounts for the appearance of the child by attributing it to a severe fright she receive by the falling down of an old white mare she was driving,” wrote Marcy. “Although I was unwilling to admit at ﬁrst that the Great Creator ever left his work in so loose a manner, that the imagination of the mother should alter or determine form or color of her children, the birth of two other albino children go further to strengthen the doctrine that the mind of the mother may affect the fetus in utero” (Marcy, 1839).

A HISTORY OF THE SCIENCE OF PIGMENTATION

John Morgan, from Philadelphia, reported the case of Adelaide, a pied Negro girl from the West Indies. Like Marcy, Morgan considered the possibility that maternal impression might cause the disorder. Morgan related that, while pregnant, the mother of the girl “. . . delighted in laying out all night in the open air, and contemplating the stars and planets. Whether the strong impression made upon the mother of Adelaide by the nightly view of the stars and planetary system may be the cause of the very extraordinary appearances in this girl, everyone will have to determine for themselves” (Morgan, 1784). During the seventeenth and eighteenth centuries, the pigment disorder that attracted most attention among the medical community, as well as the general public, was vitiligo vulgaris. In 1697, a remarkable case was presented to the Royal Society by William Byrd, a Virginian who had recently returned to London after a visit home. Under the title “An Account of a Negro-Boy that is dappeld in several Places of his Body with White Spots,” Byrd reported that the boy was born in Virginia of black parents, who was “well till 3 years old, and now was speckled of his breast and back and that no fancy had taken his Mother.” The spots he wrote are “wonderfully White, at least equal to the skin of the fairest Lady. His Spots grow continually larger and larger, and ’tis probable, if he lives, he may in time become all over white.” Thomas Jefferson described an African slave whom he encountered on his own plantation. He described the man as “A Negro man within my knowledge, born black and of black parents, developed when a boy, a white spot on his chin. This continued till he became a man, by which time it had extended over his chin, lips, and the neck on that side. He is robust and healthy and the change of color is not accompanied with any sensible disease, either general or topical” (Jefferson, 1904). Charles Peale, the painter and natural historian, also described a case under the title “An Account of a person born a Negro who afterwards became white” (Peale, 1791). The case of vitiligo that attracted most attention during this period from both lay and medical communities was Mr Henry Moss. At the age of 38 years, Moss ﬁrst noted a change in the color of his skin. The change began on his ﬁngers and hands and extended over his arms, legs, and face. Six years later, in the summer of 1796, Moss traveled from his home in Maryland to exhibit himself for money at the Black Horse Tavern in Philadelphia. He became a popular attraction and, according to one account, was so well known to the local citizens “. . . that his name was almost as familiar as John Adams, Thomas Jefferson, or James Madison” (Caldwell, 1855). Several prominent physicians from Philadelphia visited Mr Moss. Benjamin Barton, a professor at the medical school, was impressed with the distribution of the white spots. Noting that Moss’ color had disappeared completely from his armpits, Barton suggested initially that it was likely that the pigment had been washed away by perspiration. Charles Caldwell also examined Moss and was skeptical of Barton’s interpretation.

He decided to study Moss in detail. “Anxious to know as much of his case as possible, I took him in some measure under my care, . . . and . . . for a slight reward made on him such experiments as suited my purpose. While thousands visited and gazed at Moss as an object of curiosity and wonder, I alone endeavored to make him a source of scientiﬁc information.” Caldwell asked Moss “. . . to excite by exercise a copious perspiration to ascertain whether the ﬂuid perspired by the colored portions of the skin was itself colored. And I found that it was not.” Caldwell examined Moss’ skin and found that the rete mucosum had disappeared from the depigmented areas. He concluded that the rete had been removed “. . . by means of absorption.” Moss’ sensitivity to low temperatures he thought supported his theory that Moss was losing his rete. He concluded that Moss’ true skin was now “protected” from a cold atmosphere by nothing but the cuticle (Caldwell, 1855). Samuel Stanhope Smith, who had earlier pioneered the idea that African blackness was caused by environmental factors, was another visitor. He was accompanied by two other gentlemen “. . . of whom none are more capable of observing a fact of this nature with a sound and accurate judgment.” Smith paid great attention to the pattern of Moss’ pigment loss and concluded that this provided elegant proof of his hypothesis. “Although there was evidently a strong and general tendency in the constitution of this negro to a change of color, yet this tendency was much longer resisted in those parts of the body which were most exposed to the immediate action of the sun’s rays. As he was a laboring man, wherever there were rents in the thin clothes which covered him there were generally seen the largest spots of black.” From this pattern, Smith inferred that “. . . where any dark color has been contracted by the human skin, the solar inﬂuence alone, and the free contact of the external air, will be sufﬁcient to continue it a long time even in those climates which are most favorable to the fair complexion” (Smith, 1810). Benjamin Rush suggested perhaps the most unusual hypothesis to explain Moss’ pigment loss. In a paper published in 1799, Rush argued that it was the blackness of Moss, and not his white spots, that represented his disease. I shall prove, he wrote, that “. . . the color and ﬁgure of our fellow creatures who are known by the epithet of negroes are derived from a modiﬁcation of that disease which is known by the name of Leprosy.” Rush went on to explain that the black color of the negroes, their thick lips, and their insensitivity to pain were all common signs of leprosy. He suggested that it was likely that leprosy was an infectious disorder “. . . since a white woman in North Carolina living with a black husband has not only acquired a dark color but also several features of a negro.” Rush proposed that Moss was not suffering from a disease but was actually undergoing a spontaneous cure of one. “If the color of negroes is a disease . . .” he added, “. . . let science and humanity combine their efforts and endeavor to discover a remedy for it.” From his experience with other ailments, Rush suggested that bleeding or purging be tried to lessen the black color (Rush, 1799). 9

CHAPTER 1

The Beginnings of Modern Pigment Research The “early” phase of pigment research ended in the 1840s when the cell theory ended the rete’s reign as a separate amorphous layer of the skin and assigned the seat of skin color to the lower portion of a cell-rich epidermis. Microscopic anatomists had also discovered that the pigment was not in the form of a liquid or mucoid material but was rather composed of tiny intracellular granules. With the discovery that a population of dendritic cells derived from the neural crest served as the source of the coloring matter for the skin and hair, the “modern” phase of pigment research moved into high gear. For the past 150 years, researchers have relied on a melanocyte-centered paradigm to lead them along new paths of discovery. To see how far their investigations have proceeded and how wide-ranging their interests and disciplines, one needs only to look through the remaining chapters in this volume.

References Becker, S. W. Historical background of research on pigmentary diseases of the skin. J. Invest. Dermatol. 32: 185–196, 1959. Blumenbach, J. F. The Anthropological Treatises of Johann Friedrich Blumenbach. London: Longman Green, 1865. Boyle, R. Experiments and Considerations Touching Colours. London: Henry Herringman, 1664, pp. 151–167. Caldwell, C. Thoughts on the Original Unity of the Human Race. New York: E. Bliss, 1830. Caldwell, C. Autobiography of Charles Caldwell, MD. Philadelphia: Lippincott, Grambo and Co., 1855. Cruickshank, W. Experiments on the Insensible Perspiration of the Human Body. London: George Nicol, 1795. Davy, H. Essays on heat and light. In: Contributions to Physical and Medical Knowledge Principally From the West of England, T. Beddoes (ed.). Bristol: Biggs and Cottle, 1799, pp. 193–198. Gordon, J. A System of Human Anatomy. Edinburgh: William Blackwood, 1815. Harlan, R. Medical and Physical Researches. Philadelphia: Lydia Bailey, 1835. Hunter, W. Remarks on the cellular membrane and some of its diseases. Med. Observ. Inquiries 2: 26–55, 1764. Jefferson, T. Notes on Virginia, in the Writings of Thomas Jefferson. Washington, DC: Thomas Jefferson Memorial Association, 1904.

10

Josselyn, J. An Account of Two Voyages to New-England. London: G. Widdowes, 1675. Keynes, G. The Works of Sir Thomas Browne. Chicago: University of Chicago Press, 1964. Le Cat, C. N. Traite de la Couleur de la Peau Humaine en General, de celle des Negres en Particulier, et de la Metamorphose d’une de ces Coleurs en l’autre, soit de Naissance, soit Accidentellement. Amsterdam, 1765. Littre, A. La Peau in Histoire de l’Academie Royale des Sciences. Paris: Hochereau, 1720, pp. 30–32. Lovejoy, A. O. Kant and Evolution in Forerunners of Darwin. Baltimore: Johns Hopkins, 1959, pp. 173–206. Malpighi, M. De Externo Tactus Organo Anatomica Observatio. Naples: Apud Agidium Longu, 1665. Marana, G. P. Letters Written by a Turkish Spy, vol. 8. London: Vernor and Hood, 1801. Marcy, S. S. On the albino. Am. J. Med. Sci. 24: 517–518, 1839. Meirowsky, E. A critical review of pigment research in the last hundred years. Br. J. Dermatol. 52: 205–217, 1940. Montagu, M. F. A. Edward Tyson, MD, FRS, and the Rise of Human and Comparative Anatomy in England. Philadelphia: The American Philosophical Society, 1943, pp. 212–213. Morgan, J. Some account of a motley coloured, or pye, negro girl and mulatto boy. Trans. Am. Phil. Soc. 2: 392–395, 1784. Nordlund, J. J., Z. A. Abdel-Malek, R. E. Boissy, and L. A. Rheins. Pigment cell biology: An historical review. J. Invest. Dermatol. 92(Suppl.): 53s–60s, 1989. Peale, C. W. Account of a person born a negro, or a very dark mullato, who afterwards became white. Univ. Asylum Columbia Mag. 2: 409–410, 1791. Pechlin, J. N. Du Habitu & Colore Aethiopum Qui Vulgo Nigritae. Koln: Joach Reumani, 1677. Plumbe, S. A Practical Treatise on the Diseases of the Skin. London: Sherwood, Gilbert & Piper, 1837, pp. 9–16. Prichard, J. C. Researches into the Physical History of Man. London: John and Arthur Arch, 1813, pp. 163–164. Read, A. The Manual of the Anatomy or Dissection of the Body of Man. London: Francis Constable, 1642. Riolan, J. Anthropographia. Paris: Hadrianum Perier, 1618. Rush, B. Observations intended to favour a supposition that the black color (as it is called) of the negroes is derived from leprosy. Am. Philos. Soc. Trans. 4: 289–297, 1799. Schierbeek, A. (ed.). Collected Letters of Antoni van Leeuwenhoek, Vol. IV, 1683–1684, Amsterdam: Swets & Zeitlinger, 1952, pp. 245–251. Smith, S. S. An Essay on the Causes of the Variety of Complexion and Figure in the Human Species. Edinburgh: C. Elliot, 1788. Smith, S. S. An Essay on the Causes of the Variety of Complexion and Figure in the Human Species. New Brunswick: J. Simpson and Co., 1810.

The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

2

Comparative Anatomy and Physiology of Pigment Cells in Nonmammalian Tissues Joseph T. Bagnara and Jiro Matsumoto

Summary 1 The terms “chromatophore” and “pigment cell” are equivalent, although the former is usually applied to lower vertebrates and the latter to homeotherms (birds and mammals). 2 Epidermal melanophores (-cytes) are ubiquitous pigment cells present in all vertebrate classes. They are primarily elements of morphologic color change (the increase or decrease in skin pigments) and are thus responsible for the amounts of melanins found in the epidermis, feathers, and hair. 3 Dermal melanophores predominate in the dermis of lower vertebrates and are elements of both morphologic color change and physiologic color change (the rapid changes from pallor to darkening and vice versa). Other dermal chromatophores include iridophores (and leukophores) and xanthophores and erythrophores, which are also elements involved in both morphologic and physiologic color changes. The pigment composition of the three basic dermal chromatophore types is different from one another, and each contains unrelated pigments. Similarly, their cytoplasmic pigmentary organelles differ from one another in appearance, although they all seem to have a common origin from the endoplasmic reticulum. Dermal chromatophores function in background adaptation (cryptic coloration), thermal regulation, warning coloration, reproduction, and possibly in protection against solar radiation. 4 Morphologic and physiologic color changes are primarily mediated by hormones, especially melanocyte-stimulating hormone (MSH). Melatonin, melanophore-contracting hormone (MCH), catecholamines, and sex hormones play more particular roles. MSH release from the pituitary is controlled by the hypothalamus in response to background color and intensity. Melatonin is produced and released by the pineal gland (epiphysis cerebri) under conditions of darkness. While MSH causes dispersion of dermal and epidermal melanophores and aggregation of iridophores, melatonin causes aggregation of only dermal melanophores whereas epidermal melanophores and iridophores are unaffected by melatonin. Some chromatophores are directly affected by light and disperse or aggregate in response to the presence or absence of speciﬁc wavelengths of light. 5 All pigment cell types originate from the neural crest and are determined (in the embryonic sense) either in situ or by

extrinsic factors encountered during migration or after their ﬁnal destination is reached. These factors may include elements of the extracellular matrix and local factors present in the integument, including a melanization-inhibiting factor (MIF) and a melanization-stimulating factor (MSF). The last two factors may be major elements in controlling the expression of pigmentation patterns. In the zebraﬁsh model, a hostile relationship of xanthophores toward adult-type melanophores plays a key role in color pattern independent of dorsal/ventral regions. The organellogenesis of all pigmentary types seems to involve a primordial vesicle derived from the endoplasmic reticulum. 6 Such hormones as MSH, thyroid hormones, and sex hormones inﬂuence pigment cell development, but do not seem to determine pigmentation patterns. 7 The pigment cells of all vertebrate classes are homologous in that they have in common embryonic origin, pigment composition, morphology, and hormonal control. Chromatophores, usually thought to be restricted to lower vertebrates, are also present in homeotherms. Proteins or peptides that regulate pattern expression, such as MIF from amphibians and agouti protein from mammals, may be chemically and physiologically related. 8 Numerous anomalies and pathologies of lower vertebrate pigment cells exist including neoplasms such as erythrophoroma. These are not unlike malignant melanomas of higher forms. 9 As the burgeoning knowledge and technology derived from work on mammalian systems is applied to those of lower vertebrates, much can be learned about the fundamental cell biology and physiology of lower vertebrate pigmentation. In turn, because of the relative accessibility of lower vertebrates for experimentation, new knowledge about the normal and abnormal biology of mammalian pigmentation, including that of humans, will be accrued from continuing studies on these lower forms.

Historical Background For many years, knowledge, understanding, and experimentation on lower vertebrate pigmentation went on separately from that of higher vertebrates, including humans. Not until the middle of the twentieth century was it realized that the 11

CHAPTER 2

bright and often rapidly changing pigmentation of ﬁshes, frogs, and lizards (poikilotherms) is fundamentally related to the more static and more uniform pigmentation of birds and mammals (homeotherms). This divergence in understanding was signiﬁcantly registered in pigment cell terminology and nomenclature. For a detailed discussion, see Bagnara and Hadley (1973). No real confusion existed with respect to birds and mammals, in which melanin-bearing pigment cells seemed to be the only ones present and were called melanocytes without question. Among lower animals, the ﬁrst term to be applied was chromoforo (Sangiovanni, 1819; according to Parker, 1948) and, thus, with translation from the Italian, the term chromatophore has precedence. This was a term of prevalence for both lower vertebrates and invertebrates, notwithstanding the use of the sufﬁx -zyte as a substitute for -phore in much of the German literature. Thus, with respect to melanin-bearing pigment cells, the reference was to melanophores (or melanozyten). It was early realized that melanophores were present primarily in the dermis, where a variety of other brightly colored nonmelanin-bearing pigment cells were also often found, and their nomenclature provided a new source of confusion because of the discovery of speciﬁc nonmelanin pigments in or in association with these dermal pigment cells (Fox, 1953; Fox and Vevers, 1960). Thus, guanine-bearing chromatophores were often designated as guanophores based upon the pigment they contained; however, these same pigment cell types were designated by others as xantholeukophores, based upon what colors they manifested. By the end of the 1970s, much new information about structure, function, and composition of all pigment cell types had accumulated, and it was thus possible to establish a reasonable and unifying pigment cell terminology for both lower and higher vertebrates (Bagnara, 1966a). The unifying element in pigment cell terminology considered that pigment cells should be designated primarily by their appearance rather than their composition, and details of its application were presented earlier (Bagnara and Hadley, 1973). It was concluded that, among poikilotherms, there exist epidermal and dermal melanophores. The latter, together with iridophores (silvery to white) and xanthophores (yellow) and/or erythrophores (red), are also dermal. The pigmentary organelles of melanophores were referred to as melanosomes, those of iridophores as reﬂecting platelets, and those of erythrophores/xanthophores as pterinosomes and carotenoid vesicles. The embryonic origin of the various chromatophore types was addressed in early experiments performed mostly on amphibians, and on salamanders in particular. All poikilothermic chromatophores are derived from the neural crest, as was shown by the early elegant extirpation and transplantation experiments of DuShane (1935). From this site on the neural tube, chromatoblasts migrate to all regions of the integument and to other areas of the body to give rise to the particular and striking patterns of lower vertebrates. Among the early questions considered by investigators were: when were the speciﬁc chromatophores determined (in the embryonic sense); and how were the various circumscribed patterns 12

accomplished? These questions are discussed at length in an extraordinary book, Of Scientists and Salamanders, by Professor Victor T. Twitty (1966). Two of Professor Twitty’s students investigated these problems in detail. M. C. Niu (Twitty and Niu, 1954) considered the mutual inhibitory effects of individual chromatoblasts as they encountered one another during migration, while H. E. Lehman [see Lehman and Youngs (1959)] concluded that both extrinsic and intrinsic factors inﬂuenced chromatoblast expression and thus were responsible for pigmentation patterns. Later, Volpe (1964) concluded that pigmentation patterns of frogs were an autonomous function of neural crest cells, but this was disproved by Bagnara (1982) who had earlier proposed that the diverse pigment cell types of vertebrates had a common origin from a neural crest stem cell and were speciﬁed later (Bagnara et al., 1979). This concept has stood the test of time. The capacity for rapid color change (physiologic color change) among lower vertebrates has long attracted attention, and an enormous literature accumulated by the start of World War II [see Parker (1948)]. With the understanding that individual chromatophores brought about color change due to their capacity to “expand or contract,” the search was on for factors controlling these cells that allowed them to respond in an integrated fashion. Among the ﬁrst candidates for chromatophore control was the nervous system and direct innervation of chromatophores. As early as 1876, Pouchet provided information that, in teleost ﬁshes, the sectioning of peripheral nerves or the electrical stimulation of spinal nerves led, respectively, to darkening or pallor in appropriate areas. Even earlier, Brücke (1852) presented data to support the idea that color change in chameleons is under nervous control. A history of work demonstrating the control of ﬁsh chromatophores by direct innervation is well provided in Parker (1948) and Fujii (1993a). Chromatophore control by direct innervation is not a general phenomenon in lizards (Bagnara and Hadley, 1973), but such is surely the case for African chameleons (Whimster, 1971). While chromatophore control through direct innervation was shown not to be a general phenomenon among vertebrates, indirect control of chromatophores by neurotransmitter agents and hormones was shown to be signiﬁcant in most lower vertebrates (see Bagnara and Hadley, 1973; Fujii, 1993a; Parker, 1948). Of particular importance are the catecholamines, epinephrine and norepinephrine, which affect melanophores directly by causing melanophore contraction, thus leading to the phenomenon of excitement pallor seen among many ﬁshes, amphibians, and reptiles. Because these catecholamines are released by the adrenal medulla, excitement pallor can be viewed as a color change controlled by hormones. While this is the case, the role of catecholamines is minor compared with that of the hypophysial hormone, MSH. That the pituitary contains an important factor that inﬂuences pigment cells was ﬁrst realized from the simultaneous early work of P. E. Smith (1916) and B. M. Allen (1916). Early on, this factor was designated “intermedin” (Zondek and Krohn, 1932) and later it received its current designation of

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

MSH (Shizume et al., 1954). The older literature on this subject has been extensively summarized by Waring (1963) and by Bagnara and Hadley (1973). More recently, the role of MSH on poikilotherm chromatophores has been reviewed by Fujii (1993a) for ﬁshes and by Fernandez and Bagnara (1991, 1993) for amphibians. Almost 30 years ago, it became clear that MSH is the principal agent bringing about melanosome dispersion in lower vertebrates and that it does so through cyclic adenosine monophosphate (AMP) as a second messenger (Bagnara and Hadley, 1969; Goldman and Hadley, 1969). This hormone was also shown to bring about iridophore aggregation and xanthophore or erythrophore dispersion in amphibians (Bagnara, 1958, 1969). The same is now known to be true for ﬁshes (Fujii, 1993a). Before the characterization of MSH, the pituitary was sometimes viewed as possessing two opposing chromatophorotropic hormones that controlled color changes of lower vertebrates. This “bihumoral” theory of hormonal control was originally championed by Sir Lancelot Hogben in the 1940s and was reﬁned by his scholarly descendants, notably H. Waring in his monograph Color Change Mechanisms of Cold-Blooded Vertebrates (Waring, 1963). It considers that the pituitary stimulates melanophore dispersion through the elaboration of a “melanophore-dispersing hormone” (essentially MSH according to Bradshaw and Waring, 1969) and melanophore aggregation through the action of a melanophore-aggregating hormone, which had not yet been identiﬁed at that time. While the bihumoral theory failed to reach acceptance, it lurked in the background largely because of the discovery by Enami (1955) of a MCH in the pituitary of a catﬁsh. Because of his untimely death, this putative hormone received little attention until its cause was revived by Professor Bridget Baker in the 1980s. Its nature and biological signiﬁcance were reviewed by her (Baker, 1993). Among other hormones found to be of importance in the regulation of chromatophores is melatonin, a hormone produced and released by the pineal gland during darkness to bring about aggregation of dermal melanophores and thus lead to the body blanching reaction of amphibian embryos and larvae (Bagnara, 1960, 1963, 1966b). It was known from the early study of McCord and Allen (1917) that the mammalian pineal contained an agent that aggregated dermal melanophores, but it was not until the work of Lerner et al. (1958) that the agent was puriﬁed, characterized as an indole, and designated melatonin. Shortly thereafter, Bagnara (1960) announced that melatonin was elaborated from the pineal during darkness and was actually a hormone released to bring about a body blanching reaction that occurs in amphibian larvae at night in a circadian rhythm (Bagnara, 1960, 1963). Subsequently, this hormonal role for melatonin was extended to many other vertebrates including ﬁshes (see Fujii, 1993a). It is signiﬁcant that melatonin’s direct action in bringing about melanophore aggregation is restricted to dermal melanophores; thus, epidermal melanophores and dermal iridophores do not respond directly to the hormone (Bagnara, 1964a).

It has long been known that the integument of animals is directly sensitive to light in a little-understood phenomenon known as the “dermal light sense.” An excellent review of this subject has been presented by Steven (1963), who included numerous references to the possibility that pigment cells themselves are directly responsive to changes in illumination. This topic is addressed further in the later section on adaptation to darkness. Among the important concepts of pigment cell biology that have long commanded interest is that of the intracellular movements of pigment granules and, in particular, the melanosomes. Even to this day, this area of research continues to be most active and even controversial. One of the earliest theories of melanosomal movements was that of Marsland (1944), who considered that melanosomal movements were related to the solation and gelation of the chromatophore cytoplasm. Later, because it was known that colchicine causes melanosome dispersion (Wright, 1955) and reﬂecting platelet aggregation (Bagnara, 1969) in amphibians, Malawista’s (1965) conclusion that this result suggested solation of chromatophore cytoplasm seemed reasonable. Attention then shifted to piscine melanophores when Bikle et al. (1966) suggested that microtubules were important for the dispersion and aggregation of melanophores. Later, Malawista (1971) suggested a role for microﬁlaments in these processes, and thus the involvement of the cytoskeleton in melanosomal movement was established. From this period to today, most work on the translocation of pigmentary organelles has focused on ﬁsh chromatophores and especially on melanophores and erythrophores/xanthophores, and the issue has become exceedingly complicated by the many variables involved. There seem to be species differences, and there is a profound variation in the velocity of organellar movements, some occurring in seconds and others over minutes. Organellar movements involve different organellar types within the same cell, such as pterinosomes and carotenoid vesicles. In some cases, individual organellar types migrate differentially within the cell and thus preclude the possibility of passive movements during aggregation and dispersion. The actin–myosin system has been invoked in explanations of many types of organellar motility, but not all. Tubulin–dynein interactions have been considered to provide the forces that drive pigmentary organelles along the length of microtubules (Oshima and Fujii, 1987), but organellar motility can occur even in the absence of microtubules. Other force-generating proteins such as kinesin and dynamin have gained attention as possible movers of pigmentary organelles as they slide along microtubules in both aggregation and dispersion. To add to the complexity of the situation, the endoplasmic reticulum has now been invoked as a major player in organellar motility. The continuing history of this phenomenon is complicated, but discussions are available in numerous reviews that have emerged (Kimler and Taylor, 1995; Obika, 1986; Schliwa, 1984; Taylor, 1992) and in the subsequent section on molecular mechanisms for intracellular transport of pigment granules. 13

CHAPTER 2 Table 2.1. Chromatophore terminology and characterization. Chromatophore

Organelle

Pigment

Color

Source

Epidermal melanophore (-cyte) Melanophore

Melanosome Melanosome Pterino-melanosome

Eumelanin Eumelanin Pterorhodin–eumelanin

Black, brown Black, brown Black, red

Iridophore

Reﬂecting platelet (iridosome)

Structural colors and iridescence

Leukophore

Leukosome (refractosome) Pterinosome (xanthosome) Carotenoid vesicles Pterinosome (erythrosome) Carotenoid vesicles Cyanosome

Purines, especially guanine; some pteridine crystals Uric acid (?), other purines (?) Pteridines

Fishes, amphibians, reptiles Fishes, amphibians, reptiles Phyllomedusine frogs, rhacophorid frogs (?) Fishes, amphibians, reptiles

Structural white

Fishes

Yellow to orange

Fishes, amphibians, reptiles

Carotenoids Pteridines

Orange to red

Fishes, amphibians, reptiles

Carotenoids ?

Blue

Mandarin ﬁsh

Xanthophore

Erythrophore

Cyanophore

Epidermal melanin unit: an association of an epidermal melanophore (-cyte) with adjacent keratinocytes that serve to receive cytocrine melanin. Dermal chromatophore unit: an association of the dermal chromatophores (xanthophores or erythrophores, iridophores, and melanophores) to bring about integrated color changes. These are especially important in the expression of green colors in amphibians and reptiles.

Current Concepts It is impossible in one chapter alone to consider all the salient aspects of lower vertebrate pigment cells that should be of interest and importance to scholars of pigmentation. Thus, in this section, an attempt is made to focus on those topics that may be most valuable to current workers in the ﬁeld. These topics are considered to be perhaps most relevant to the ongoing research of today. At the same time, it should be understood that much of the knowledge presented in the previous section on history remains current. Much of it will not be discussed further because, unfortunately, current relevance dictates the direction of current research support, even at the expense of allowing important concepts to lie fallow. In recent years, most pigmentary research on poikilotherms has centered on ﬁshes and amphibians (Bagnara, 1976). The cytophysiology of ﬁsh chromatophores has been the subject of detailed review (Fujii, 1993a, b). Accordingly, amphibian examples will be stressed here.

Chromatophore Characterization Because of new knowledge gained during the past few years, it is time to re-examine chromatophore terminology and perhaps establish a nomenclature that can be used universally. Table 2.1 presents a suggested terminology to be used for the near future. It does not differ markedly from that which we presented earlier (Bagnara, 1966a; Bagnara and Hadley, 1973); however, other terms that have emerged more recently are included as are some lesser known synonyms that are indicated in parentheses. Figures 2.1–2.3 demonstrate some of the brightly colored pigment cells. 14

Fig. 2.1. Living red-backed salamander, Plethodon cinereus, showing the distribution of erythrophores and melanophores over capillaries between the skin glands of the dorsal surface. A few silvery iridophores are also visible (see also Plate 2.1, pp. 494–495). [From Bagnara, J. T. Color change. In: Physiology of the Amphibia, vol. 3, B. Lofts (ed.). New York: Academic Press, 1976, pp. 1–52 with permission.]

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

Fig. 2.2. A living scale from the red-colored variant of swordtail ﬁsh, Xiphophorus helleri. Note the presence of numerous erythro(xantho)phores with red pterinosomes in the periphery and yellow carotenoids in the center under a dispersed state (see also Plate 2.2, pp. 494–495). [From Bagnara, J. T., and M. E. Hadley. Chromatophores and Color Change: The Comparative Physiology of Animal Pigmentation. Englewood Cliffs, NJ: Prentice-Hall, 1973, with permission.]

Melanophores and Melanocytes Whether these cells should be designated -phores or -cytes is a question that has often been addressed, and there seems to be no answer other than to let use by the scientiﬁc community be the ultimate judge. In an earlier attempt to reconcile the problem, Fitzpatrick et al. (1966) suggested that “melanophore” be the term of preference for lower vertebrates because this pigment cell was important in rapid color change phenomena (physiologic color change) and involved the rather rapid and profound intracellular movements of melanosomes, the melanin-containing organelles. It was suggested that the term “melanocyte” be used for higher vertebrates primarily because their epidermal pigment cells were of a form different from that of dermal melanophores and presumably were not involved in physiologic color change. Actually, some of the rationale for their premise no longer holds, but their conclusion that melanocyte be used for birds and mammals seems justiﬁed because this term has come to be used almost exclusively for melanin-bearing pigment cells by current workers. Accordingly, although this chapter is devoted to the pigment cells of lower vertebrates, the mutuality of nomenclature for melanin-bearing pigment cells will be acknowledged by liberal use of the sufﬁx -cyte, as in melanophore (-cyte). Epidermal Melanophores (-cytes) While the epidermal melanocytes of birds and mammals had long been recognized and reasonably characterized, it was not until the work of Hadley and Quevedo (1967) that it was generally realized that this very same pigment cell type exists in the epidermis of lower vertebrates where it functions as it does in homeotherms. Indeed, it has been observed to be a prominent pigment cell of most lower vertebrates, from primitive ﬁshes to reptiles, where it exhibits its typical spindle shape (Fig. 2.4) and

Fig. 2.3. Melanophore and xanthophores from the teleost, Fundulus heteroclitus. Note that the xanthophores appear as aggregated oil droplets (see also Plate 2.3, pp. 494–495). [From Bagnara, J. T., and M. E. Hadley. Chromatophores and Color Change: The Comparative Physiology of Animal Pigmentation. Englewood Cliffs, NJ: Prentice-Hall, 1973, with permission.]

Fig. 2.4. Epidermal melanophores of adult Rana pipiens, dispersed at left and aggregated at right. Note cytocrine melanin in adjacent epidermal cells. Each epidermal melanocyte with its surrounding epidermal cells comprises an epidermal melanin unit (courtesy of Professor M. E. Hadley).

15

CHAPTER 2

V pre

mel

V mel pre

Fig. 2.6. Dispersed (left) and aggregated (right) dermal melanophores in the tail ﬁn of Xenopus larvae.

Fig. 2.5. Dermal melanophores from larvae of Pachymedusa dacnicolor. Left, melanosomes (mel) and ﬁbrillar premelanosomes (pre) are typical of early stages. Right, at metamorphic climax, melanosomes begin to develop into the adult form and, at this stage, are characterized by elevation of the limiting membrane and the rough appearance of the eumelanin mass; prominent Golgi (G) and numerous vesicles (V) are found within the cytoplasm together with premelanosomes (pre) (from Bagnara et al., 1978b).

serves to deposit considerable quantities of cytocrine melanin into adjacent epidermal cells (Bagnara and Hadley, 1973). Each epidermal melanophore (-cyte), together with the epidermal cells that it serves, comprise a true “epidermal melanin unit” as ﬁrst described by Fitzpatrick and Breathnach (1963). Much of what is known about organellogenesis of lower vertebrate epidermal melanophores is extrapolated from studies on mammals (see Chapter 7); however, there are probably few differences among all vertebrate groups with respect to melanosome formation. The melanin-containing organelles of poikilotherms are similar in shape and size to those of mammals and include ﬁbrillar premelanosome stages (Fig. 2.5). The similarity between epidermal melanophores among the various taxonomic groups is also registered in their responses to hormones. Epidermal melanophores of lower vertebrates respond to MSH by dispersing their melanosomes just as do mammalian epidermal melanocytes in the presence of the hormone (Snell, 1967). This response to MSH is rather interesting in view of the fact that their thin spindle shape precludes much of a role in physiologic color change. In contrast, epidermal melanophores of lower vertebrates play an important role in morphologic color change, as is manifested by the signiﬁcant amounts of melanin that they deposit in the epidermis (see Figs 2.4 and 2.8). Obviously, there is an increase in melanogenesis in these cells in response to MSH stimulation, but it is not known how this occurs. Presumably, MSH affects melanogenesis as it does in mammals (see Chapter 19). In contrast to dermal melanophores, epidermal melanophores are not sensitive to melatonin. They do not aggregate in response to melatonin in any vertebrate studied so far (Bagnara and Hadley, 1970a). 16

Fig. 2.7. Dispersed dermal melanophores from the dorsal surface of a Xenopus larva demonstrating the broad ﬂattened form of this dermal pigment cell.

Dermal Melanophores These prominent black or brown chromatophores of lower vertebrates are profoundly important not only because they provide most of the dark pigmentation of ﬁshes, amphibians, and reptiles, but because they are exceedingly active in physiologic color change in these animal groups (Fig. 2.6). They are large cells with broad dendrites; they usually reside in the upper portion of the dermis, just below the basal lamina. Often, they are ﬂat and thus occupy a broad area of the dermal surface (Fig. 2.7). This is especially true of organisms in larval or embryonic stages. Flat melanophores seem more prevalent when the dermis is relatively thin. In some adults, especially those of amphibians and reptiles, which possess a thicker dermis, the form of dermal melanophores is more threedimensional. Such forms often possess “dermal chromatophore units,” to be discussed subsequently, composed of an upper layer of xanthophores (yellow pigment cells) lying

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

EM X I

M

Fig. 2.8. Transverse section of the dorsal skin from an MSH-treated adult R. pipiens showing basket-shaped dermal melanophore (M). Iridophores are in a punctate state (I), and the clear xanthophore layer (X) is also visible. An epidermal melanin unit is seen in the epidermis with an epidermal melanophore (EM) and surrounding epidermal cells containing cytocrine melanin (from Bagnara and Hadley, 1969).

just below the basal lamina and above a layer of iridophores, and a basal layer of melanophores. These dermal melanophores extend processes upward to terminate between the xanthophores and iridophores (reﬂecting cells) in amphibians (Bagnara et al., 1968) or above the xanthophores in lizards (Taylor and Hadley, 1970). In both cases, the form of the melanophore becomes basket-like (Fig. 2.8). Melanophores of the dermal type are not restricted to the dermis, but are often present on and in internal organs such as the liver, kidney, heart, thymus, and gonads, on blood vessels, in the peritoneum, and on the meninges (see Figs 2.25, 2.26, and 2.30). That these are of the dermal type is inferred from their sensitivity to hormones such as MSH or melatonin. For example, in hypophysectomized larvae of Xenopus, these deep melanophores are aggregated (see Fig. 2.26), in contrast to those of intact larvae, which are dispersed. Similarly, such deep melanophores aggregate in the presence of melatonin (see Fig. 2.34). Melanosomes of dermal melanophores are not unlike those of epidermal melanophores except for the fact that they are often larger than those of the latter. Generally, they are more spherical in form and about 0.5 mm in diameter. Most poikilotherm dermal melanosomes, especially those of amphibians, possess typical ﬁbrillar premelanosomes (Fig. 2.5) and form in the usual way from the Golgi–endoplasmic reticulum (GERL). In contrast, piscine melanosomes, such as those of the goldﬁsh, form in a different manner (Turner et al., 1975). Here, a multivesicular body of endoplasmic reticular origin forms the premelanosome and no ﬁbrillar substructure is observed. Whereas melanosomes of all vertebrates seem consistently similar, there are remarkably unusual types found in some taxonomic groups. The most notable of these are the melanosomes of the phyllomedusine frogs (leaf frogs) of the New World (Bagnara et al., 1973; Bagnara, 2003). Dermal

Fig. 2.9. Wholemount of portion of brown dorsal skin of P. dacnicolor that had been masked and photographed with epiillumination. The sharp transition between the masked (above) and unmasked (below) skin is evident. The preparation involved passage through solvents, thus the iridophore surfaces are exposed because of the loss of yellow pigments and blue coloration is expressed. Note the red color of the melanophore pigment (see also Plate 2.4, pp. 494–495).

melanophores of adults of these species are a red color (Fig. 2.9) and, as shown in Figure 2.10, they may contain large ﬁbrillar melanosomes (Taylor and Bagnara, 1969). They may or may not contain eumelanin, but their predominant pigment is pterorhodin, a pteridine dimer (Misuraca et al., 1977). They are unquestionably melanophores on the basis of their form, position, and function (Figs 2.9 and 2.11); moreover, in at least the Mexican leaf frog, Pachymedusa dacnicolor, adult melanosomes are derived from typical melanin-containing melanosomes (Bagnara et al., 1978a). Although these unusual organelles have never been named formally, we have designated them as pterino-melanosomes in Table 2.1. At metamorphic climax, the typical larval melanosomes undergo a “redifferentiation” such that pterorhodin ﬁbers begin to deposit upon the eumelanin core of the melanosome to form a compound organelle containing a central kernel of eumelanin surrounded by a concentric mass of pterorhodin (Bagnara and Ferris, 1974). It is likely that such unusual organelles are not restricted to the phyllomedusine frogs (Bagnara and Ferris, 1975), but may be present in taxonomically unrelated frogs such as the rhacophorids (J. T. Bagnara, personal observations; P. Schwalm, personal communication). Iridophores and Leukophores Pigment cells that function primarily through the reﬂection of light from the surface of orderly distributed organelles, or reﬂecting platelets, are called iridophores (Fig. 2.12) (Bagnara, 1966a; Taylor, 1969). Although not true pigments, the purines guanine, uric acid, hypoxanthine, and adenine have long been considered to be deposited in crystalline form in the reﬂecting platelets, which are usually arranged in oriented stacks. More recently, studies of pure iridophores from frog larvae grown 17

CHAPTER 2

A

0.1 µ

B

0.1 µ

Fig. 2.10. Unusual large melanosomes in dermal melanophores of two phyllomedusine species, P. dacnicolor (A) and Agalychnis callidryas (B). The melanosomes of the two species are each larger than the typical vertebrate melanosome and contain the pigment pterorhodin. Each melanosome is composed of a ﬁbrous mass enclosed by a limiting membrane, but that of A. callidryas lacks the electron-dense eumelanin kernel typical of the P. dacnicolor melanosome (from Bagnara et al., 1973).

Fig. 2.11. An adult Pachymedusa dacnicolor in a “semi-natural” outdoor cage in full illumination on a light background. The black plastic letter masks, UA (University of Arizona), to the left and right of this brown-adapted frog were removed after having been in place on the frog’s back for 1 hour. Note the green color that developed under the mask (see also Plate 2.5, pp. 494–495). [From Iga, T., and J. T. Bagnara. An analysis of color change phenomena in the leaf frog, Agalychnis dacnicolor. J. Exp. Zool. 192:331–342, 1975.]

in cell culture have revealed that guanine was the sole purine present; moreover, it was discovered that some pteridine pigments were also present in these cells (Bagnara et al., 1988). It seems that reﬂecting platelets may contain both crystalline purines and pteridines. The orientation of these stacks of platelets determines the nature of the pigmentary function of the iridophore. Normally, when viewed with reﬂected light, iridophores appear to contribute a metallic gold or silver luster (see Figs 2.1 and 2.47–2.49). When viewed with transmitted light, iridophores exhibit blues, greens, and reds. These are structural colors probably arising from thin-layer interference and the scatter or the diffraction of light from the stacks of reﬂecting platelets. Some iridophores examined with reﬂected light appear to be blue, while others appear to be brown or tan in color. These colors probably relate to the size, shape, orientation, and conformation of reﬂecting platelets. A pene18

trating analysis of the optics of iridophores in ﬁshes has been presented by Fujii (1993a), and a more speciﬁc study of the structural colors of spiny lizards has been made by Morrison (1995). The latter study emphasizes the role of thin-layer interference in establishing the various iridescent colors in these lizards. A complete analysis and understanding of the structural colors of any individual iridophore or group of iridophores is very difﬁcult for numerous reasons, not least of which is the size and shape of reﬂecting platelets. Among the more common types found in amphibians and ﬁshes are those represented by long, thin proﬁles (Fig. 2.12). Among the leaf frogs, small bead-like or cubic reﬂecting platelets prevail (Fig. 2.13) as they often do in some lizards (Fig. 2.14). Distances between reﬂecting platelets are very important, and only slight changes in spacing can markedly alter the

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

ians and reptiles (see Bagnara and Hadley, 1973; Bagnara et al., 1978b). An elementary scheme of how these colors are brought about is shown in Figure 2.17.

Fig. 2.12. An iridophore from the dorsum of an adult R. pipiens showing stacks of reﬂecting platelets (actually spaces previously occupied by purine crystals that shattered and dissolved during sectioning and staining). An individual reﬂecting platelet is trapped between two lobes of the nucleus. Scale = 1 mm (from Bagnara, 1976).

apparent color, as was shown for the tree frog Hyla arborea by Nielsen (1978). Another chromatophore active in structural coloration has been recognized in ﬁshes (see Fujii, 1993a). This chromatophore, the leukophore, seems to be somewhat different from the iridophore; however, it functions much as iridophores and probably utilizes purines as pigments in its pigmentary organelles, as do the reﬂecting platelets of iridophores. The introduction of another chromatophore designation may add to nomenclatural confusion; however, out of deference to recent investigators of piscine chromatophores who have described these cells, the leukophore is acknowledged as a distinctive chromatophore (Menter et al., 1979). Very likely, it is a particular type of iridophore found primarily in ﬁshes or in anomalous situations in other vertebrates, for example, as in the periodic albino mutant of Xenopus (Fukuzawa, 2004). With the acceptance of the existence of both iridophores and leukophores, new names for their inclusions (Table 2.1) must be recognized; these include “leukosome” or “refractosome.” Despite the existing differences between iridophores and leukophores, their similarities are such that it warrants their being grouped together as a basic pigment cell type. Iridophores play an important role in imparting blue coloration as a structural color (Figs 2.15 and 2.16). Blue coloration is relatively uncommon among most poikilotherms and blue pigments are rare; however, blue as a structural color is important because it plays a major role in the elaboration of green coloration, which is very common among amphib-

Xanthophores and Erythrophores Dermal chromatophores that range in color from pale yellow to bright red are designated xanthophores (yellow) or erythrophores (red) depending upon how yellow or red they appear (Figs 2.1–2.3). Despite their differences in color, xanthophores and erythrophores should be considered as the same basic pigment cell type (Table 2.1). At the beginning of the twentieth century, they were called lipophores because many of these cells contain fat-soluble yellow, orange, or red carotenoid pigments that are localized in fatty vesicles (Bagnara, 1966a; Bagnara and Hadley, 1973). It was not until much later (Bagnara, 1961; Bagnara and Obika, 1965; Obika, 1963; Obika and Bagnara, 1964) that another group of pigments, pteridine in nature, was found to be even more important pigments of xanthophores and erythrophores. Soon afterwards, it was discovered by Matsumoto (1965a) that pteridine pigments are localized in organelles called pterinosomes and that both carotenoid vesicles and pterinosomes are often found within the same cell, be it a xanthophore or an erythrophore (Figs 2.2, 2.13, and 2.18). The speciﬁc color manifested by the cell often depends upon the pattern of carotenoids and pteridines it contains. In addition, it is possible that there exist other pigments so far undiscovered. All these facts were important considerations in the decision to designate chromatophores in accordance with their color rather than their chemical composition (Bagnara, 1966a). Although the association of carotenoid pigments with xanthophores or erythrophores has a long history, there have been relatively few deﬁnitive studies on their speciﬁc carotenoid content. Moreover, it has been only recently that attempts have been made to correlate carotenoid composition with xanthophore or erythrophore ultrastructure. Matsui et al. (2002) have observed that the bright red ventral integument of the Japanese newt, Cynops pyrrogaster, contains beta-carotene and at least ﬁve other carotenoids including canthaxanthin and lutein. They suggest that these pigments are contained in either thin-walled vesicles or thicker-walled “ring-type” vesicles. The precise localization of carotenoids in these vesicles remains unclear. In a general sense, perhaps the most important of the xanthophore and erythrophore pigments is the pteridine group. Pteridines are a group of yellow, orange, or red compounds having the structure of pyrimido[4,5-b]pyradine and are known to be synthesized from guanosine triphosphate (GTP) by GTP-cyclohydrolase I (GTP-CHI) that has a variety of isoforms (Thony et al., 2000). The vast majority of naturally occurring pteridines have the structure of 2-amino-4oxopteridine and are called unconjugated pteridines collectively. This terminology is convenient to discriminate them from folic acid or its related compounds, which have a long side-chain at position 6 of the pteridine ring. Thus far in medical research, much attention has been focused on 19

PT

CV

RP

MS

5 Fig. 2.13. Transverse section through the dorsal dermis of P. dacnicolor. Xanthophores are ﬁlled with pterinosomes (PT) and carotenoid vesicles (CV). Iridophores contain reﬂecting platelets with round or oval proﬁles, which seem to be strung together in a bead-like fashion. Melanosomes (MS) are typical for this species. The organelle indicated by an arrow may be a pterinosome. Magniﬁcation ¥ 6000 (from Taylor and Bagnara, 1972).

Fig. 2.14. Iridophores of the lizard, Urosaurus ornatus, at low magniﬁcation at left (¥ 2000) and high magniﬁcation at right (¥ 75 000). Note the rectangular proﬁles of reﬂecting platelets and their orderly arrangement. The image on the left includes views of melanophores (bottom) and xanthophores (top) (courtesy of Dr Randall Morrison).

20

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

Fig. 2.15. Melanophores (aggregated) and iridophores (dispersed) in a wholemount dorsal skin preparation of Rana pipiens photographed with transmitted light. Note the varied structural colors evident in the iridophores (see also Plate 2.6, pp. 494–495). [From Bagnara, J. T., and M. E. Hadley. Chromatophores and Color Change: The Comparative Physiology of Animal Pigmentation. Englewood Cliffs, NJ: Prentice-Hall, 1973, with permission.]

biopterin, particularly its tetrahydro form, because of its role as a cofactor for hydroxylation of phenylalanine and tryptophan (Nicol et al., 1985). In lower vertebrates, large amounts of unconjugated pteridines accumulate speciﬁcally in xanthophores and erythrophores. Some, such as sepiapterin, serve as yellow pigments by themselves, whereas others, such as biopterin, 2-amino-4-oxopterin, 7-oxobiopterin (ichthyopterin), and isoxanthopterin, are colorless in nature. The roles of these colorless pteridines are unknown, but it is possible that they function in the absorption of ultraviolet light as their spectrophotometric characteristics include strong absorption in the 340–360 nm region (Mori et al., 1960; Tschesche and Glaser, 1958). It is also known that some unconjugated colorless pteridines, such as drosopterins, form dimers that act as chromophores (Pﬂeiderer, 2001). Biosynthetic pathways of pteridines common to lower vertebrates are well established, as summarized in Figure 2.19

Fig. 2.16. Wholemount of portion of brown dorsal skin of P. dacnicolor that had been masked and photographed with epiillumination. The sharp transition between the masked (above) and unmasked (below) skin is evident. The preparation involved passage through solvents, thus the iridophore surfaces are exposed because of the loss of yellow pigments and blue coloration is expressed. Note the red color of the melanophore pigment (see also Plate 2.7, pp. 494–495).

(Masada et al., 1990; Ziegler et al., 2000). As seen in this scheme, it is apparent that pteridine metabolism is regulated by three key enzymes including GTP-CHI, 6-pyruvoyl-tetrahydropterin synthase (PTPS), and sepiapterin reductase (SPR) (Masuda et al., 1990; Ziegler et al., 2000). Therefore, mutations in genes associated with these enzymes, particularly GTP-CHI, would inevitably affect development and differentiation of xanthophores and thus yield pigmentary mutants. This seems to be the case in zebraﬁsh where there exists a variety of mutants, three of which are associated with xanthophore pigmentation and pteridine composition, including one that displays an alteration in pteridine metabolism similar to a mutation in Drosophila (Odenthal et al., 1996). The autonomy of xanthophores and erythrophores for pteridine biosynthesis was shown by elegant techniques of classic embryology. Obika (1963) demonstrated by in vitro 21

CHAPTER 2

Fig. 2.17. Diagrammatic interpretation for the basis of green coloration in amphibians and other vertebrates. As light strikes the surface of an animal such as a frog, short wavelengths of light (blue–violet) are largely absorbed by the ﬁltering xanthophore or yellow pigment layer; the rest are scattered by the iridophore or scattering layer. Long wavelengths (red–orange) largely pass through the ﬁltering and scattering layers of the skin and are absorbed by the melanophore or melanin layer. Intermediate wavelengths (yellow–green) pass through the ﬁltering layer and are scattered from the surface of the iridophore layer (Tyndall scattering) and pass back through the ﬁltering layer. Thus, the light reﬂected from the surface contains a high proportion of yellow–green wavelengths and the animal appears green (from Bagnara and Hadley, 1973).

Fig. 2.18. Schematic interpretation of the dermal chromatophore unit based upon observations from several anurans. Adaptation to a dark background is represented. Processes from basally located melanophores, between overlying xanthophores and iridophores, become ﬁlled with melanosomes and thus obscure light reﬂection from the iridophore surface (from Bagnara et al., 1968).

hanging drop culture of urodele (Triturus and Hynobius) neural crest that yellow pigmentation of larval xanthophores occurs simultaneously with formation of pteridines in simple physiological salt (Holtfreter) solution. Richards and Bagnara (1967) further demonstrated autonomous formation of species-speciﬁc pteridines, such as 6-hydroxymethylpteridine, by larval urodele xanthophores through the use of xenoplastic transplantation of neural crest between Axolotl and Pleurodeles. These experiments also revealed the autonomous expression of “Pleurodeles blue,” a pteridine relative and a presumed yellow pigment in larval xanthophores. This novel 22

pyridone N-glycoside (Yoshida et al., 1988) seems to be newt speciﬁc (J. T. Bagnara, unpublished). Combined cytochemistry and chemical analytical studies on skin preparations provided proof that pteridines function as pigments in these brightly colored pigment cells. Their intracellular localization in particular cytoorganelles, as in the aforementioned pterinosomes, was ascertained by ultracentrifugal fractionation of subcellular components of xanthophores and erythrophores in sucrose density gradients (Matsumoto, 1965a; Matsumoto and Obika, 1968; Obika and Matsumoto, 1968). Skin samples laden solely with these pigment cells were collected from ﬁshes or frogs by careful dissection of homogeneously colored spots or stripes. Combined pteridine assay and electron microscopy on separated fractions disclosed that the vast majority of pteridines were detected in a fraction composed of large cytoplasmic granules, the pterinosomes, although certain amounts were present in fractions composed of membranous elements, mostly those of smooth endoplasmic reticula or Golgi complexes. Carotenoids, when present, were detectable in fractions composed of small vesicles or membrane fragments, indicating an apparent disparate distribution of pteridines and carotenoids within the cells. The morphology of pterinosomes is characterized by a spherical shape, slightly larger than melanosomes, and a distinct boundary of unit membrane enclosing an internal structure consisting of thin concentric lamellae or deposits of thin ﬁbrils (Matsumoto, 1965a; Matsumoto and Obika, 1968; Obika and Matsumoto, 1968). Little is known about the relationship between the internal structure of pterinosomes and the composition of pteridines deposited. In developing xanthophores or erythrophores, multivesicular bodies are present

GTP GTP-CHI O

OH OH N

HN H2N

N

PHO

N H

H

O

C

C CH2O

P O

H

H

OH

3

H

Dihydroneopterin-3-phosphate PTPS

O HN H2N

N

O

OH OH N

C

C CH2OH

H

H

HN H2N

N

N

H N

HO O C C CH3 H N H H

H2N

N

H HH O N C C CH3 OH H N H H

HN H 2N

6-Lactoyltetrahydropterin

N

N H

H

Dihydropterin

SPR O

H HO H N C C CH3 H OH N H H

H H

H2N

XOR

AR O

N HN

6-Pyruvoyltetrahydropterin

Neopterin

HN

O

N

6-(1’-Hydroxy, 2’-oxopropyl) tetrahydropterin

O

O

H N

HN

O

H

N

HN

H H2N

N H

N

H

H2N

Dihydroxanthopterin

N

N

Pterin

SPR

XOR SPR

N

HN H2N

N

O

O H

O

N H

C C CH3 OH H H

H2N

N

O

O

O HN

H2N

N

N

H2N

Xanthopterin

N

N

H

N H

O

Isoxanthopterin

H H H C C CH3 OH OH H N H H N

HN HN

O

H N

HN

Tetrahydrobiopterin

Sepiapterin

SPR

H H H H N C C CH3 OHOH N HH H

HN

N

Quinoiddihydrobiopterin

O N

HN H2N

N

N H

H H C C CH3 OH OH H H

Dihydrobiopterin

O N

HN

COOH

O N

HN

H H C C CH3

O N

HN

OHOH H2N

N

N

Pterin-6-carboxylic acid

H 2N

N

N

Biopterin

H 2N

N

N H

H H C C CH3 OHOH O

Ichthyopterin

Fig. 2.19. Biosynthetic pathways of unconjugated pteridines common to xanthophores and their related pigment cells of lower vertebrates. Abbreviations of enzymes: AR, aldose reductase; GTP-CHI, GTP-cyclohydrolase I; PHO, phosphatase; PTPS, 6-pyruvoyl-tetrahydropterin synthase; SPR, sepiapterin reductase; XOR, xanthine oxidoreductase (adapted from Katoh and Akino, 1986; Masada et al., 1990; Takigawa and Nakagoshi, 1994; Ziegler et al., 2000).

23

CHAPTER 2

in the vicinity of Golgi complexes or GERL (J. Matsumoto et al., unpublished data). The similarity of these multivesicular bodies to Stage I melanosomes suggests that pterinosome genesis shares an origin common to that of melanosomes. These observations are consistent with the concept of the common origin of pigment cells (Bagnara et al., 1979). Cyanophores A most remarkable discovery was made a decade ago by Goda and Fujii (1995) who found blue dermal chromatophores that they designated cyanophores. These chromatophores contain a true blue pigment of unknown chemical nature. The blue pigment is found in ﬁbrous organelles that they termed cyanosomes. Whereas cyanophores have been found, so far, in only two species of callionymid ﬁsh, it may be premature to afford them legitimate status. However, they are real and unique and must be recognized. Accordingly, they are included in Table 2.1.

Fig. 2.20. Green eggs (yolk plug stage) of P. dacnicolor. Eggs normally vary in color from blue-green to yellow-green (see also Plate 2.8, pp. 494–495).

Other Unknown Chromatophores Among some of the less studied groups of amphibians, there exist unusual pigmentary organelles in chromatophores that may be xanthophores or erythrophores. Schwalm and McNulty (1980) described some unusual chromatophores that seem to contain large vesicles with pigments that have not been elucidated. The dendrobatid (poison arrow) frogs also contain brightly colored chromatophores with vesicular organelles, the compositions of which are also unresolved (J. T. Bagnara, personal observations).

The Amphibian Egg as a Pigment Cell One would hardly think that there could be any relationship between the pigmentation of the integument and that of the eggs. On the contrary, the pigmentation of the eggs of frogs and skin have much in common both by homology and by analogy (Bagnara, 1985). The frog integumental pigmentary pattern of a dark dorsum and light ventrum is paralleled in the animal and vegetal poles, respectively, of frog eggs. Moreover, melanin is the dark pigment in both cases, and the pigment in each is contained in melanosomes. It appears that this pattern of pigmentation serves protective functions in both cases. In addition to dark pigmentation, the bright colors of the integument have much in common with the comparable colors that are found in the eggs of some species. In particular, the eggs of leaf frogs, the Pyllomedusinae, which are usually laid on green vegetation above the surface of pools of water, are often bright green (Fig. 2.20). A fundamental difference in egg pigmentation is that the green color is arrived at in a manner different from that of the skin. In both cases, the green color derives from a combination of blue and yellow; however, the blue element of the skin is a structural color whereas that of eggs is provided by the blue pigment, biliverdin IXa. This can be seen in Fig. 2.21, in which a blue area from a portion of P. dacnicolor skin is manifested when the overlying yellow pigment is leached from normally green skin. In contrast, as shown in Fig. 2.21, a blue pigment 24

Fig. 2.21. Pigments extracted from P. dacnicolor eggs. Left, biliverdin IXa; right, lutein (see also Plate 2.9, pp. 494–495).

(biliverdin) and a yellow pigment (lutein) provide the green color of P. dacnicolor eggs (Marinetti and Bagnara, 1982, 1983). In the absence of yellow pigments from xanthophores in the integument, frogs become blue. In Fig. 2.22, a blue individual of P. dacnicolor is shown. This frog was apparently deﬁcient in xanthine dehydrogenase, a key enzyme in pteridine biosynthesis, and thus failed to produce pteridine pigments (Frost, 1978; Frost and Bagnara, 1978). In cases of carotenoid deﬁciency, the skin of this species also becomes bluish. This is a consequence of the absence of yellow carotenoid pigments from the xanthophores. In Fig. 2.23, sibling juveniles of P. dacnicolor are shown, one green and the other blue. The latter was raised on carotenoid-deﬁcient crickets (potato-fed) and the former on carrot-fed crickets (J. T. Bagnara and G. V. Marinetti, unpublished). Analysis of the crickets revealed a preponderance of carotenes in the carrot-fed individuals and very low levels of any carotenoid in crickets fed potatoes. Of special interest here is the fact that the integumental

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

Fig. 2.22. “Blue Boy,” a young male P. dacnicolor, which lacked xanthophore pigments from his metamorphosis until sexual maturity when he gradually began to turn green. Presumably, he was xanthine dehydrogenase-deﬁcient, but eventually produced enough pteridine pigments to restore normal green coloration (see also Plate 2.10, pp. 494–495).

Fig. 2.23. Sibling froglets of P. dacnicolor. The green frog was raised on crickets fed carrots exclusively, while the blue frog was fed potato-raised crickets that were carotenoid-deﬁcient (see also Plate 2.11, pp. 494–495).

carotenoids of P. dacnicolor appear to be exclusively acidic xanthophylls. Thus, it appears that the frog liver must undertake major metabolic steps to convert carotenes to xanthophylls. How this occurs in frogs is unknown. Moreover, the transport of the xanthophylls from the liver to the skin is a mystery. Probably, this transport to the integument is similar to that of lutein, an acidic xanthophyll, to the egg wherein vitellogenin seems to serve as a transport protein for both biliverdin and lutein (Bagnara, 1985). Such transport in insects, that of biliverdin and lutein, involves speciﬁc transport proteins (see Kanost et al., 1990; Kawooya et al., 1985) and thus, by analogy, it seems reasonable to consider that the same may hold for amphibians.

Chromatophore Function A major function of integumental chromatophores is in background adaptation (cryptic coloration). Most lower vertebrates exhibit rapid body color changes for use in camouﬂage or background matching, as is well exempliﬁed by the tropical ﬂounder (Ramachandran et al., 1996). The simplest type, adaptation to black or white backgrounds, mainly involves the participation of both melanophores and iridophores. Thus, on black surfaces, melanosomes within melanophores become widely dispersed within the dendritic processes of the cell. At the same time, iridophores are minimally visible as their reﬂecting platelets aggregate to the center of the cell. In other words, black coloration is maximized and white is minimized. 25

CHAPTER 2

Fig. 2.24. In vitro response of R. pipiens skin photographed with reﬂected light. Left, in Ringer solution, iridophores, which appear silvery, are expanded; dermal melanophores are punctate. Right, in the presence of caffeine, intracellular levels of cyclic AMP become elevated; thus, iridophores are contracted to the punctate state while melanophores are well expanded (from Bagnara, 1976).

On white surfaces, these two pigment cells respond in an opposite manner. Adaptation to more particular backgrounds, such as to speciﬁc colors or shades or even mottled surfaces, may entail the participation of all three basic pigment cell types. Aside from their role in cryptic coloration, chromatophores function in a variety of other roles. Thermal regulation, protection from solar radiation, nuptial coloration, and warning coloration represent a few of the other more prominent roles of lower vertebrate chromatophores. Because of their roles in so many adaptive activities, the physiology of lower vertebrate chromatophores is by necessity exceedingly complex and is an important component of the integrative biology of poikilotherms. A fundamental feature is the fact that color change results from the translocation of pigmentary organelles within the cell (Figs 2.4 and 2.6). In early times, chromatophores were referred to as being “expanded or contracted” with the implication that the cell boundaries were actually motile. We now know that pigment cells are ﬁrmly attached to the extracellular matrix and that contractile motility actually refers to movements of the pigmentcontaining organelles within the chromatophore. Nevertheless, unfettered chromatophores, as in cell culture, do have the capacity to expand and contract, but it is not known whether this is a passive result of intracellular organellar motility or the result of some other contractile event. In any case, we now 26

consider color change to result from the aggregation or dispersion of pigmentary organelles within the cell (Fig. 2.24). This type of chromatophore motility has been the subject of continued scrutiny in recent years, especially as new knowledge about the cytoskeletal system of pigment cells has accumulated. The responses of pigment cells during color change are integrated, and mechanisms for this integration reside in several signaling mechanisms. As color change often results from the visual perception of background change, the nervous system has long been considered to be a key element in chromatophore signaling, and direct innervation of chromatophores was an early focus of investigation. Over the years, we have learned that chromatophore control through direct innervation occurs in some poikilotherms, but is by no means a general phenomenon. While best evidence for such innervation comes from studies of piscine chromatophores (see Bagnara and Hadley, 1973; Fujii, 1993a), not all orders of the class Osteichthyes possess innervated chromatophores. Early reports of possible innervation of amphibian chromatophores have been discounted (Iga and Bagnara, 1975); however, the early reports of the late 1880s describing innervation of reptilian chromatophores (chameleons) certainly remain valid (Whimster, 1971). In other lizards, such as Anolis, direct nervous control of chromatophores does not

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

exist (see Bagnara and Hadley, 1973); thus, we are left with the unanswered question of how pervasive direct innervation of chromatophores exists among reptiles.

Responses of Pigment Cells Physiologic Color Change Physiologic color changes are deﬁned as those rapid changes in color caused by an intracellular movement of pigmentbearing organelles (Parker, 1948). As has been indicated, the response can be elicited by a variety of stimuli such as light or hormone action. The response is rapid, requiring a few minutes or a few hours to occur, and is transitory, i.e. the animal either regains its original coloration or assumes an intermediate state depending on stimulatory cues. Perhaps the most common manifestation of physiologic color change occurs in background adaptation. On a darkcolored background, amphibians that can adapt contain melanophores with dispersed melanosomes, while those on light-colored backgrounds are pale because their melanosomes have aggregated to a perinuclear position within the cell (Figs 2.4, 2.6 and 2.24). Iridophores respond in an opposite manner, being dispersed on light-colored backgrounds and aggregated on dark backgrounds. Apparently, the lateral eyes play a fundamental role in governing these background responses of both melanophores and iridophores. In most amphibian larvae, there is a distinct point in development at which they become able to adapt to background. Before this time, when they are not able to adapt to background, they remain dark colored whether on a light or a dark background, and they are said to be in a primary phase or stage. Older larvae that can adapt to background are said to be in a secondary stage. The secondary stage describes what essentially constitutes adaptation to background based upon the reception of light through the eyes and the control of MSH release from the pituitary. Melanosome aggregation can occur in primary-stage larvae in darkness, but this does not result from alterations in MSH release, but relates to other physiological systems that operate in darkness as mentioned earlier and to be described more fully later. It should be noted that onset of the secondary phase varies with species: whereas melanophores of Xenopus larvae respond to background changes from very young stages onward, larvae of Rana and Hyla do not display the secondary response until they are several weeks old, and some Bufo larvae never seem to acquire the secondary phase. These observations of the ontogeny of physiologic color change have been made primarily on amphibians and, although much is known about the physiology of chromatophores of ﬁshes, the majority of the work has been on adults or at least on post-embryonic stages. Morphologic Color Change Color changes that are evoked slowly and result from alterations in the amount of pigment contained in the integument have long been called morphologic color change (Parker, 1948). Morphologic color change is a relatively slow process that includes the synthesis or destruction of relatively large

amounts of pigment in response to a persistent stimulus. Background adaptation is a common cause of morphologic color change: animals maintained on dark backgrounds develop more melanin whereas those on light backgrounds lose their melanin (Dawes, 1941). The opposite is true of iridophores with respect to their purine pigments (Bagnara and Neidleman, 1958; Fernandez, 1988; Fernandez and Collins, 1988; Taylor, 1969). The total increase or decrease in the amount of pigment can result from several causes, one of which relates to the number of chromatophores present. For example, Xenopus larvae that have been deprived of their pituitary gland (and hence have no melanophore stimulation) contain many fewer dermal melanophores than they do normally (Bagnara, 1957). That the hypophysis exerts its inﬂuence on morphologic color change by causing a profound proliferation of dermal melanophores has been elegantly shown (Fig. 2.25) by Pehlemann (1967a, b, 1972). Probably, the increase in chromatophore number following persistent stimulation also results from the synthesis of pigments in hitherto undifferentiated melanoblasts. This phenomenon seems to be a general one and provides a valid interpretation of the fact that iridophores appear in the tail ﬁn of Xenopus larvae that have been deprived of their hypophysis (Fig. 2.26), although such chromatophores are not observed in intact normal larvae (Bagnara, 1957). Amphibian iridophores are markedly inhibited by MSH, both physiologically and morphologically (Bagnara, 1958). In his PhD dissertation, Hadley (1966) described how, in juvenile Xenopus maintained on dark backgrounds for 10 weeks, there is a ﬁvefold increase in the number of web melanophores present in comparison with controls maintained on white backgrounds. The spacing of these new melanophores seems to exclude the presence of doublets as would be expected in cases of rapid increase in melanophore numbers due to proliferation; thus, it is presumed that these new melanophores were derived from latent melanoblasts that underwent melanization in response to the elevated blood levels of MSH present in those animals on black backgrounds. Similar results were obtained by Fernandez and Bagnara (1991, 1993), who studied morphologic color change in leopard frogs (Figs 2.27 and 2.28). They showed that the intense melanization corresponded directly to elevated blood levels of MSH as measured by radioimmunoassay (RIA) (Fig. 2.29). Deﬁnitive proof that latent undifferentiated melanoblasts are the source of new melanophores comes from an unpublished study described by Bagnara and Fernandez (1993) in which the pars intermedia of juvenile Xenopus were implanted into the lower unpigmented jaw of larvae (Fig. 2.30). Melanophores differentiated in an area concentric to the implant, implying that they were derived from latent melanoblasts. Manifestations of morphologic color change occur at the organellar level and in some cases can be seen with the electron microscope. For example, after administration of large dosages of MSH to adult frogs, a condensation of the reﬂecting platelets can be seen in iridophores (Bagnara et al., 1968, 1969; Taylor, 1969). Through action on the adjacent epidermal cells, epidermal 27

CHAPTER 2

Fig. 2.25. Details of the upper right side of the head of a Xenopus larva, (A) 1, (B) 5, (C) 7 days after transection of the hypophysial stalk leading to greatly increased levels of circulating MSH and melanophore proliferation. A single melanophore (arrow) can be traced through two mitotic divisions in (B) and (C) (courtesy of Dr F. W. Pehlemann; from Pehlemann, 1967b).

melanophores also play an important role in morphologic color change (Hadley and Quevedo, 1967). During profound melanophore stimulation, such as prolonged exposure to a dark background, pronounced cytocrine activity of the dendrites of epidermal melanophores causes melanin to be deposited in adjacent epidermal cells (Figs 2.4 and 2.8). This process in frogs has not been studied with the electron microscope, and the mechanism of melanin deposition is not understood; however, studies made at lower magniﬁcation indicate that considerable amounts of melanin are deposited in the epidermis in this manner. In adult frogs, cytocrine deposition of melanin is the primary means of morphologic color change (Hadley and Quevedo, 1967). The importance of epidermal melanophores (-cytes) in homeotherms is well known, and it has been suggested that epidermal melanocytes of all vertebrates are homologous in that they seem to be identical in every way (Bagnara and Ferris, 1971).

The Control of Color Change The Pituitary and the Role of MSH Fig. 2.26. A Xenopus larva hypophysectomized as a tailbud embryo. Note the presence of iridophores and the great reduction in melanophores of the integument, brain case, and other deep areas (from Bagnara, 1957).

28

The ability of lower vertebrates to carry out morphologic and physiologic color change in response to appropriate environmental cues is based upon the existence of precise control mechanisms involving the pars intermedia of the pituitary and the release of MSH. Whereas the existence of this hormone was inferred from experiments on poikilotherms, characteri-

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

α–MSH (pg/ml)

200 175 150 125 100 75 50 25 0

α–MSH (pg/ml)

Fig. 2.27. Dorsal (top) and ventral (bottom) views of adult R. chiricahuensis adapted to dark backgrounds (left individuals) or white backgrounds (right individuals). Note the profound darkening of the dark background-adapted individuals and especially the ventral surface (from Bagnara and Fernandez, 1993).

1800 1600 1400 1200 1000 800 600 400 200 0

Rana pipiens

White

Gray

Fig. 2.28. Dorsal (top) and ventral (bottom) views of adult R. pipiens adapted to dark backgrounds (left individuals) or white background (right individuals). Note the darkening of the dorsal surface of the dark background-adapted individuals and the lack of darkening of the ventral surface of the dark background-adapted individuals (from Bagnara and Fernandez, 1993).

Black

Rana chiricahuensis

Gray Black White Background color

Fig. 2.29. Plasma a-MSH of R. pipiens and R. chiricahuensis on white, gray, and black backgrounds. Each bar represents the mean (±SE) of 1–4 frogs. Plasma a-MSH of frogs on a black background is greater than those on white or gray (P < 005). Note the particularly profound increase in plasma a-MSH of R. chiricahuensis (modiﬁed from Fernandez and Bagnara, 1991).

zation of MSH was achieved from mammalian sources. A personal account of the biochemical characterization of the MSHs in the mid-1950s has been graciously written by Professor Aaron B. Lerner (1993). It was not until some 30 years later that poikilothermic MSHs were isolated, puriﬁed, and characterized from ﬁshes and amphibians. Thus, some idea of

Fig. 2.30. Lower jaw of a larva of Xenopus laevis that had been implanted subcutaneously with the pars intermedia of a froglet. A spherical blood clot marks the site of the transplant, which releases MSH chronically. Note that the nearby dermis is populated with melanophores distributed over the graft. No melanophores were observed in the dermis of the jaw before the transplantation (from Bagnara and Fernandez, 1993).

the phylogeny of MSH was obtained (Dores et al., 1993). It has been shown that, although the tridecapeptide a-MSH is a ubiquitously secreted peptide derived from the ancestral molecule, proopiomelanocortin, N-acetylation of the hormone is 29

CHAPTER 2

a key element in the phylogeny of the molecule. Like that of mammals, ranid a-MSH is N-acetylated, a scheme inherited from an ancient tetrapod. During evolution, the scheme was retained except among some of the squamate reptiles, which have lost this capacity. Among the latter is the fence lizard Anolis, which secretes a non-N-acetylated a-MSH that is nevertheless a functional melanotropin.

Melanophore Stimulation The most well-known activity of MSH is to stimulate the dispersion of melanosomes within melanophores of poikilotherms. In the absence of MSH, melanophores assume a punctate (contracted) conﬁguration in which melanosomes are concentrated in the central (perinuclear) portion of the melanophore (Figs 2.4, 2.6 and 2.24). Both dermal and epidermal melanophores respond to stimulation by MSH by demonstrating physiologic color change, whereas prolonged exposure to the hormone, especially at high concentration, is conducive to morphologic color change. It has been thought that morphologic color change is an indirect function of MSH activity that actually results from persistent physiologic color change; however, Pehlemann’s (1972) demonstration of increased melanization resulting from enhanced mitotic activity in melanophores under MSH stimulation indicates that some expressions of morphologic color change are independent of physiologic color change. It is concluded that MSH affects both dermal and epidermal melanophores and is capable of eliciting morphologic color change in both chromatophores. The possible role of MSH on pigment pattern formation through its effects on morphologic color change will be considered later. Iridophore Effects Earlier work on color change centered around melanophore responses. During later years, however, attention was given to the effect of MSH on iridophores, the reﬂecting pigment cells commonly found in poikilotherms (Figs 2.8, 2.31, and 2.32). In normal larvae and in intact adult frogs as well as in ﬁshes, iridophores are in a punctate state due to a concentration of reﬂecting platelets toward the center of the cell (see Bagnara and Hadley, 1973; Fujii, 1993a). In the absence of the pituitary, reﬂecting platelets are dispersed and the iridophore exists in an expanded state. The “silvery” appearance of hypophysectomized ranid larvae observed by Smith (1916) and Allen (1916) results from the “expansion” of iridophores and the “contraction” of melanophores; in intact larvae, the reverse is true. Smith (1920) recognized that the hypophysis exerted an inﬂuence on iridophores, which he referred to as xantholeukophores; however, except for Hogben and Winton (1924) and Stoppani et al. (1954), most other workers emphasized melanophore effects, and gradually the prominent role of iridophores in amphibian color change was overlooked and almost forgotten. Interest in amphibian iridophores was renewed when it was discovered that these cells are controlled by MSH just as are melanophores (Bagnara, 1958). Substantial proof for this 30

Fig. 2.31. A mixed primary cell culture of chromatophores, containing mostly iridophores, isolated from R. pipiens tadpoles. Left, attached iridophores in their normal semi-aggregated (expanded) state. Right, a similar culture after 1 h of exposure to [Nle4,D-Phe7]a-MSH (10-5 M). Both photographs were taken under epi-illumination. Note the aggregated (contracted) state of the treated iridophores, which have actually detached from the surface of the culture dish.

Fig. 2.32. Cells similar to those in Fig. 2.31, viewed in thin sections. Upper left, a rounded iridophore before attachment. Bottom, a dispersed iridophore attached to the culture dish. Upper right, detached and rounded iridophores following MSH treatment.

concept was obtained when highly puriﬁed a- and b-MSH ﬁrst became available and were shown to cause iridophore aggregation in both hypophysectomized larvae (Bagnara, 1964b) and in isolated skins of adult frogs (Hadley, 1966; Hadley and Bagnara, 1969). That iridophores are not only sensitive to MSH, but also react by aggregating rather than dispersing under its inﬂuence led to the question of whether both melanophore expansion and iridophore contraction require the same sites on the MSH molecule (Fig. 2.24). That this is indeed the case was demonstrated through the utilization of the fact that, in adult frog skins, the entire MSH molecule is not essential for melanophore-expanding activity (Lee et al., 1963) and that some melanophore response can be induced

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

by the centrally located polypeptide sequence, His-Phe-ArgTrp-Gly. When pentapeptide and larger synthetic MSH peptides were injected into hypophysectomized larvae, deﬁnite iridophore aggregation was observed, and the iridophore response was always paralleled by melanophore dispersion (Bagnara, 1964b). A similar parallelism of iridophore and melanophore responses was obtained with a thermal polymer of arginine, glutamic acid, histidine, phenylalanine, and tryptophan (Bagnara and Hadley, 1970b). This early work that utilized short amino acid sequences of the MSH molecule to study iridophore responses made little impact because it was very much obscured by the rapid accumulation of new knowledge about MSH chemistry that was mounting up. This work centered on the frog skin bioassay, which was generally considered to involve melanophore responses, although Hadley and Bagnara (1969) clearly described the integrated responses of both melanophores and iridophores in this assay. Early on, Bagnara (1958) had demonstrated that the alkali-enhanced activity of MSH peptides was evidenced in both melanophore and iridophore responses. In later years during the development of superpotent a-MSH analogs (Sawyer et al., 1980), their effects on iridophores were not observed. Recently, such a study has been carried out (J. T. Bagnara et al., unpublished) on tadpole iridophores in culture and, as can be seen in Figure 2.31, iridophores exposed to the superpotent analog [Nle4,D-Phe7]a-MSH indeed become aggregated. This is of special interest not only because it demonstrates iridophore sensitivity to the hormone, but because the response displays both aggregation of reﬂecting platelets and actual contraction of the iridophore, causing it to round up and detach from its substrate (Fig. 2.32). Early work on the role of MSH in iridophore control was done on amphibians, and only in recent years was it shown that piscine iridophores respond to the hormone in the same way (see Fujii, 1993a). Leukophores, however, may provide a bit of a paradox in that leukophores of the medaka may disperse in the presence of MSH (Negishi and Obika, 1980). Iridophores play a role in both physiologic and morphologic color changes. When high doses of MSH are administered or when hormone treatment is prolonged, larvae of some species such as Rana sylvatica display a morphological response so great that the many iridophores of hypophysectomized larvae may completely lose their pigment (Bagnara, 1958; Bagnara and Neidleman, 1958). This morphological effect of MSH on iridophores has been studied by Taylor (1969), who points out that reﬂecting platelets in iridophores become thinner in frogs receiving MSH injections. The diminution in reﬂecting platelet thickness is a morphological manifestation of a quantitative loss of measured purines. Xanthophore Effects Relatively little is known about the physiological responses of amphibian xanthophores or erythrophores to MSH because these chromatophores are difﬁcult to see. They are often pale, especially at their margins, and the presence of large numbers of iridophores and melanophores serves to mask the xan-

thophores. Nevertheless, there are at least some clear cases in which xanthophores are inﬂuenced by MSH. In adults of Hyla arenicolor, xanthophores of skins kept in Ringer solution are aggregated; however, upon administration of MSH, these cells expand (Bagnara, 1969; Bagnara et al., 1968). In more recent times, there have been numerous studies on the xanthophores and erythrophores of ﬁsh, and it is now well documented that both these chromatophores generally disperse under the inﬂuence of MSH (see Fujii, 1993a). The morphological effect of MSH on xanthophores of amphibians is more prominent and is best demonstrated by quantitative changes in pteridine pigments. In the skin of hypophysectomized larvae, the content of pteridines is considerably lower than that of intact larvae, but it returns to normal when such larvae are injected with MSH (Bagnara, 1961, 1969; Bagnara and Neidleman, 1958). Xanthophores in normal larvae of the salamander Pleurodeles waltlii are expanded such that their broad arms form a continuous yellow sheet over the surface. Xanthophores of hypophysectomized larvae are also dispersed; however, individual processes are thin and delicate and individual chromatophores stand out clearly. Thus, the morphological effect of MSH on xanthophores is manifested by the amount of pteridine pigments contained and is reﬂected in the general appearance of the chromatophore. In Pleurodeles, there is also a marked difference in the total carotenoid content between normal and hypophysectomized larvae (Bagnara, 1969). It is not known, however, whether the diminution of carotenoids from the skin of hypophysectomized larvae is a result of the absence of MSH or an indirect manifestation of the lack of a hypophysis. Despite the profound effect of MSH on the pteridine content of xanthophores, there has been no observation of an MSH effect on the pterinosome. No differences have been noted between the pterinosomes of skins from normal frogs and those from frogs receiving MSH injections. Cyanophore Effects With their discovery of cyanophores in both the mandarin ﬁsh, Synchiropus splendidus, and the psychedelic ﬁsh, S. picturatus, Goda and Fujii (1995) showed that these chromatophores responded to various stimulatory cues. Whereas their exposure to norepinephrine resulted in slow aggregation, a-MSH induced pigment dispersion.

Melanin-Concentrating Hormone (MCH) MCH is a cyclic heptadecapeptide secreted from the posterior lobe of the pituitary that has the capacity to bring about melanin aggregation in piscine melanophores (Kawauchi et al., 1983). It seems to be present primarily in teleost ﬁshes (Sherbrooke et al., 1988). This neuropeptide has been found in the brain of other lower vertebrates where its neuronal cell bodies seem to be localized in the hypothalamus (Baker, 1993). In most species, only a few of its axons project to the posterior pituitary; however, in teleosts, the neurohypophysial lobe receives many MCH ﬁbers, and thus the neuropeptide has 31

CHAPTER 2

achieved legitimate hormonal status in this group. It seems to function in the regulation of skin color through its ability to bring about pigment aggregation within melanophores and apparently in xanthophores and erythrophores as well (Baker, 1993). It is possible that the use of MCH in the control of color change in this group of ﬁshes is an evolutionary novelty that arose near the end of the Paleozoic or during the early Mesozoic, just before or in the evolution of the Holostei, a group ancestral to modern teleosts (Sherbrooke and Hadley, 1988). Baker (1993) has summarized the evidence that MCH is used in color change in teleosts. Among the points that she raises is the fact that MCH can induce melanin concentration in isolated ﬁsh skin melanophores. The differential effects observed following the use of MCH fragments on different species suggests variation in the MCH receptor. MCH has the capacity to antagonize the melanin-dispersing action of aMSH directly at the melanophore level. MCH may also play a role in pigmentary control by depressing release of MSH from the pituitary. Among the fruits of continuing investigation in this ﬁeld is the revelation by Oshima and Wannitikul (1996) that cyclic AMP is probably the messenger for MCH action.

The Effects of Other Hormones on Pigment Cells In addition to MSH, other hormones are known to affect chromatophores. Two of these, melatonin and epinephrine, will be discussed in the following sections because they are associated with speciﬁc responses. The thyroid has long been implicated in pigmentary changes that occur during amphibian metamorphosis. Woronzowa (1932) reported that thyroid extracts affected the spotting pattern in metamorphosing ambystomids, and Kollros and Kaltenbach (1952) observed pigmentary changes in the vicinity of thyroid implants in Rana pipiens larvae. A profound increase in chromatophore number occurs at metamorphic climax in such larvae (Bagnara and Hadley, 1973). Similarly, speciﬁc pteridine changes were induced in localized areas of the skin of Pleurodeles larvae following the implantation of thyroxine cholesterol pellets (Bagnara, 1964c). Among pteridine-associated changes is the abrupt disappearance of “Pleurodeles blue,” long thought to be a pteridine, but now known to be a novel pyridine derivative (Yoshida et al., 1988). The mode of action of thyroxine on the metabolism of this compound is a question that begs an answer. Effects of thyroxine on physiologic color change have been observed. Chang (1957) indicated that the blanching of frogs following administration of thyroxine may be attributable to an inhibition of MSH release from the pars intermedia. The same may hold for salmonids (Bagnara and Hadley, 1973). The direct action of this hormone on pigment cells was found in the in vitro study by Wright and Lerner (1960). Triiodothyronine is more potent than thyroxine or any other known hormonal agent, except for melatonin, in reversing the action of MSH on isolated frog skins (Bagnara and Hadley, 1970a; Hadley and Bagnara, 1969). 32

Studies on other vertebrates have long indicated that steroid hormones inﬂuence pigmentation, and such an effect on amphibian pigmentation was reported by Himes and Hadley (1971), who observed that progesterone exerts an MSH-like response on frog skins in vitro. A strong case for a role for sex hormones in in vivo color changes was presented by Richards (1982), who demonstrated a profound alteration of chromatophore expression by sex hormones in the Kenyan reed frog Hyperolius. Adaptive Mechanisms Background Adaptations MSH has long been considered the major hormone involved in background adaptation, and this peptide has provided the basis for the unihumoral theory of chromatophore control (see Parker, 1948). This theory considers that, during adaptation to white backgrounds, little or no MSH is released from the pars intermedia, resulting in low circulating levels of this hormone and a consequent lack of stimulation of chromatophores. Thus, melanophore pigments are aggregated and iridophore pigments are dispersed. During black background adaptation, MSH is released leading to a dispersion of melanophore pigments and an aggregation of reﬂecting platelets within iridophores; consequently, the animal darkens. This concept is supported by observations of cytological changes in the pars intermedia of frogs in correlation with background adaptation (Cohen, 1967; Imai, 1971; Perryman, 1974; Saland, 1967). The appearance of pars intermedia cells in frogs adapted to black backgrounds is consistent with a state of hormone synthesis and release. This was also the conclusion of Burgers et al. (1963) made on the basis of actual MSH assays. In background adaptation, it is implicit that the animal can perceive differences in background. It seems that the lateral eyes are involved as blinded animals equilibrate to an intermediate state of pigmentation and do not alter coloration in response to changes in background (see Parker, 1948; Bagnara and Hadley, 1973). Moreover, direct electrophysiological studies by Dawson and Ralph (1971) have shown that changes in illumination of the lateral eyes of adult R. pipiens are clearly recorded in the pars intermedia. The mechanism by which the retina discriminates background difference is not understood, although it has long been thought (Butcher, 1938) that dorsal and ventral regions of the retina have different sensitivities to incident and reﬂected light and thus allow appropriate perception of albedo (the amount of light reﬂected from the substrate, as explained in Fig. 2.33). Ultimately, background adaptation is regulated by the control of MSH release through the involvement of the hypothalamus. The control is mediated through the inhibition of MSH release, as was suggested by Etkin (1941, 1962a, b) who along with others obtained extremely dark tadpoles after isolating the pituitary by hypophysial transplantation or by hypophysial stalk section (Fig. 2.25). During the past 30 years, an immense literature has accumulated in this area, and the current state of knowledge has been summarized by René et al. (1993) and Roubos et al. (1993). The evidence is over-

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

Light source

Reflecting substrate High albedo

Absorbing substrate Low albedo A

Light source

High albedo environment

Light source

Low albedo environment

Fig. 2.33. Perception of light by Arizona tiger salamanders differs between white and black substrates. Light is reﬂected from a white substrate and provides a high albedo, whereas a black substrate absorbs light and the albedo is low. Larvae in a turbid pond (lower left) experience a high albedo because of light reﬂected from suspended particles, while those in a clear pond are subjected to a low albedo because light is absorbed by plants and the dark-colored pond bottom (courtesy of P. Fernandez; from Fernandez and Collins, 1988).

whelming that hypothalamic inhibition of MSH secretion is controlled by catecholamines, notably dopamine. Presumably, the pars intermedia chronically releases MSH, and this release is controlled through neurosecretory ﬁbers that reach the pars intermedia from the hypothalamus. Perception of a light background (high albedo) results in inhibition of MSH release and consequent paling of the animal. On a dark background (low albedo), hypothalamic neurosecretory neuron release of dopamine is reduced and resulting high levels of MSH lead to darkening (Figs 2.27–2.29). Adaptation to Darkness Unlike adults, larval stages of most amphibians display a

B

Fig. 2.34. Xenopus larvae, stage 42. (A) Larva under normal room illumination on a white background; larva does not background adapt because it is in a primary stage. (B) Similar larva in darkness for 1 h; note prominent melanophore contraction in response to melatonin release from the pineal (from Bagnara, 1976).

remarkable ability to blanch when they are maintained in darkness (see Bagnara, 1965, 1966b; Bagnara and Hadley, 1970a). This blanching reaction was ﬁrst observed long ago (Babak, 1910); however, it was only much later that a basic mechanism controlling this response was described (Bagnara, 1960, 1961). The proposed mechanism describes a role for the pineal in a normal physiological function. It is essentially hypothetical, but it remains unchallenged and is widely accepted (Bagnara and Hadley, 1970a; Bogenschütz, 1965). The hypothesis suggests that, under conditions of darkness, the pineal is stimulated to release melatonin, presumably a pineal hormone, into the general circulation. Melatonin exerts a profound contracting effect on dermal melanophores leading to a rapid blanching (Fig. 2.34). As ﬁrst described by Bagnara (1960), the involvement of the pineal relates to two aspects of its physiology: light reception and endocrine function. The former has been a role attributed to the pineal since before the classic work of von Frisch (1911) on ﬁshes and has since found substantial support from both ultrastructural and electrophysiological studies (see Eakin, 1973). The role of the pineal as an endocrine organ is more obscure and requires a fuller explanation. The ﬁrst evidence that the pineal contains a humoral agent is attributable to the studies of McCord and Allen (1917), who discovered that feeding mammalian pineals to tadpoles evoked a profound blanching. Later, Lerner et al. (1958) isolated a potent melanophore-contracting agent from beef pineal glands, which they identiﬁed as melatonin (Nacetyl-5-methoxytryptamine). A detailed analysis of the data supporting the concept that the body-blanching reaction is mediated by the pineal has been presented elsewhere (Bagnara, 1965; Bagnara and Hadley, 1970a; Eakin, 1973); however, a few important points need to be made here. First of all, it should be mentioned that 33

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blinded larvae become pale when they are placed in darkness (Bagnara, 1960; Laurens, 1915, 1917). Moreover, the blanching reaction is abolished by “pinealectomy” (Bagnara, 1960, 1963; Charlton, 1966). Temporal events in the onset and recovery from the blanching reaction are consistent with the view of an endocrine mediation of the response. Of many indoles tested, only melatonin is a potent melanophorecontracting agent (Quay, 1968; Quay and Bagnara, 1964), and the pigmentary changes induced by the action of melatonin duplicate the responses that occur in darkness. Young tadpoles, not yet able to respond to dark backgrounds by inhibiting MSH release from the pars intermedia, are in the primary stage and thus have dispersed melanophores. Such larvae blanch when placed in darkness due to the direct effects of melatonin on dermal melanophores causing them to aggregate. In larvae that possess epidermal melanophores or iridophores, these chromatophores do not change in darkness because they do not respond directly to melatonin. While these data support the hypothesis that the bodyblanching reaction of amphibian larvae is controlled by the pineal, it must be emphasized that this mechanism is restricted to larvae and does not appear to be generally functional in adults. It is well known that adult amphibians do not blanch in darkness (see Parker, 1948). Moreover, both epidermal and dermal melanophores in skins of adult frogs are generally unresponsive to the administration of melatonin, as has been demonstrated both in vitro (Bagnara and Hadley, 1970a; Hadley and Bagnara, 1969) and in vivo. Whether they lack the melatonin receptor originally deduced by Heward and Hadley (1975) on the basis of structure–function studies and characterized recently by expression cloning from Xenopus (Ebisawa et al., 1994) is not known. Among the various color changes displayed by vertebrates, those resulting from the direct action of light are among the most striking. Perhaps the most remarkable of these is the tail-darkening reaction observed when hypophysectomized Xenopus larvae were placed in the dark (Bagnara, 1957) and subsequently shown to occur in isolated tails maintained in darkness (Fig. 2.35) (Bagnara, 1957, 1966b; Burgers and van Oordt, 1962; van der Lek et al., 1958). Under usual conditions of illumination, dermal melanophores of the ﬁn are punctate, or nearly so, and the tail is essentially transparent. When isolated tails or whole tadpoles are placed in darkness for half an hour or more, a profound dispersion of melanosomes occurs in these melanophores so that the tail becomes black. Upon resumption of illumination, these melanophores quickly revert to the original punctate state and the tail becomes pale. Illumination of given regions of the tail (Bagnara, 1957) or of individual melanophore processes (van der Lek, 1967) can cause aggregation of melanosomes, suggesting that the response is mediated by a photochemical system operating at the level of the chromatophores themselves. The tail-darkening reaction was ﬁrst considered to be a rather peculiar specialization of Xenopus larvae, but we now know that it occurs in larvae of a Mexican leaf frog, P. dacnicolor (Bagnara, 1974), and in many other hylid tadpoles. 34

Fig. 2.35. In vitro tail-darkening response of isolated tails from Xenopus larvae under normal conditions of illumination (top) and in darkness (bottom). The tail-darkening response was prevented in a central area which was left illuminated (center) (from Bagnara, 1957).

The dynamic light-sensitive tail ﬁn melanophores have been used to address another infrequently considered phenomenon of chromatophore physiology, that of the energy requirements for the aggregation–dispersion phenomenon. Some investigators have considered that pigment aggregation is the principal energy-requiring step (Horowitz, 1958), whereas others have concluded that dispersion is the principal energy-demanding process. Iga and Bagnara (1975) have demonstrated that both aggregation and dispersion of melanosomes are energy requiring and will not occur in the absence of oxygen (Fig. 2.36). Furthermore, they showed that, although darkness-induced darkening of Xenopus tail ﬁn melanophores does not occur in oxygen-free media, restoration of oxygen to such tails even after they have been returned to lighted conditions leads to a dispersion aftereffect. This indicates that the phenomena of light reception and melanosome dispersion are two separate components of the tail-darkening reaction. In addition to the tail-darkening reaction, the direct effect of light on melanophores cultivated in vitro has been reported. Kuhlemann (1960) indicated that embryonic melanophores of Xenopus grown in tissue culture respond to illumination by becoming punctate (conﬁrmed more recently by Daniolos et al., 1990), just as do embryonic melanophores of neural crest explants of several anurans (Fig. 2.37) (Bagnara and Obika, 1967). The signiﬁcance of these direct effects of light on melanophores grown in culture is not known.

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES A

Fig. 2.37. Neural crest explant of Xenopus. Under normal room illumination, melanophores remain in an aggregated state (left) but, following a 45-min period in darkness, these melanophores become punctate (right) (modiﬁed from Bagnara and Obika, 1967). B

C

Fig. 2.36. (A) Response of tail ﬁn melanophores of a Xenopus tadpole to darkness (D) and to light (L) in Ringer solution; the dark period is indicated on the abscissa by a solid black bar. Melanosome dispersion takes place gradually in darkness, and aggregation occurs quickly in light. (B) Inhibition of melanosome dispersion in the tail ﬁn melanophores of a Xenopus tadpole in oxygen-free Ringer solution and the dispersion “aftereffect,” which occurs in oxygen-containing Ringer solution. The dark period is indicated on the abscissa by a solid black bar. (C) A typical example of retardation of melanosome aggregation in the tail ﬁn melanophores of a Xenopus tadpole in oxygen-free Ringer solution; the dark period is indicated on the abscissa by the solid black bar (D). Pigment dispersion of the melanophores was induced by dark treatment of the tails for 45 min in Ringer solution, and then the tails were transferred into oxygen-free Ringer solution in a dark room (from Iga and Bagnara, 1976).

In addition to amphibian melanophores, examples of direct light sensitivity of ﬁsh chromatophores are well documented. Iga and Takabatake (1986) have demonstrated that teleost melanophores may respond to local light stimulation by aggregating in darkness and dispersing in the light. Of special interest among ﬁshes is the demonstration of light-sensitive iridophores in the neon tetra (Lythgoe et al., 1984; Nagaishi and Oshima, 1989), such that the distance between reﬂecting platelets within these cells is changed through the action of the light. In a search for the basis of this photosensitivity of iridophores, Lythgoe et al. (1984) demonstrated the presence of an opsin-based visual pigment in these iridophores, either or both rhodopsin or/and porphyropsin. The presence of visual pigments in a photosensitive chromatophore is both fascinating and signiﬁcant and asks the question whether similar visual pigments are present in other chromatophores such as the light-sensitive melanophores discussed previously. It seems likely that, as more species are studied with respect to lightinduced color changes, more unusual phenomena will be revealed. Such seems to be the case with P. dacnicolor, in which the skin of brown individuals displays a direct photosensitivity (Iga and Bagnara, 1975). When masks are placed on the brown surface, the skin beneath becomes green in exact conformation with the mask (Fig. 2.11).

Mechanisms of Hormone Action on Pigment Cells The Cyclic AMP as a Second Messenger of MSH Action In accordance with the ﬁrst messenger–second messenger scheme (Sutherland et al., 1965), MSH acts as a ﬁrst messenger and brings about its effects by promoting an intracellular increase in a second messenger, 3¢,5¢-cyclic adenosine monophosphate (cyclic AMP), which is in turn responsible for the particular response of the effector cell (Sutherland et al., 1968), in this case pigment cells. Thus, cyclic AMP mimics the action of MSH by darkening frog skin in vitro (Bagnara and Hadley, 1969; Bitensky and Burstein, 1965; Novales and Davis, 1967). The response is not as effective as that of MSH itself or as that of the dibutyryl derivative of cyclic AMP (Goldman and Hadley, 1969); however, the response to either of these cyclic nucleotides is truly MSH-like, for cytological 35

CHAPTER 2

examination of darkened skin reveals that both melanophores and iridophores react. Probably, both iridophores and melanophores contain similar or identical MSH receptor sites, and the different events in the two cells following MSH stimulation could then be attributable to the interaction of the second messenger and the speciﬁc distal functional elements of the particular chromatophore. Further support for the involvement of the ﬁrst messenger–second messenger concept in the regulation of chromatophore control is derived from the fact that methylxanthines, such as caffeine or theophylline, can bring about both iridophore and melanophore responses (Fig. 2.24). Methylxanthines are known to increase cellular levels of cyclic AMP by inhibition of cyclic nucleotide phosphodiesterase (Butcher and Sutherland, 1962; Sutherland and Rall, 1958). Furthermore, it has been reported that MSH stimulates cyclic AMP formation in frog skin in correlation with the degree of darkening (Abe et al., 1969). The parallelism between iridophore and melanophore responses to the various MSH peptides, be they unnatural synthetic partial sequences or entire molecules such as a-MSH or b-MSH (Bagnara, 1958, 1964b), is to be expected from the point of function. The most efﬁcient darkening response involves both melanosome dispersion and reﬂecting platelet aggregation. Accordingly, it seems logical to conclude that, despite the differing intracellular responses of these two divergent chromatophores, they must each possess similar MSH receptors. In modern times, the quest for characterization of the MSH receptor has led to the discovery of several different MSH receptors that function in accordance with speciﬁc roles for MSH (Cone et al., 1993). At present, at least ﬁve receptors are known from mammals for the melanocortin [MSH/adrenocorticotropic hormone (ACTH)] peptides. These melanocortins are labeled numerically in order of their discovery, with melanocortin-1 (MCR-1) assigned to the surface of melanocytes. It is known through the use of various speciﬁc a-MSH antagonists that frog melanophores possess at least one MCR, but which of the ﬁve is not known (Hruby et al., 1995). An interesting question concerns the possible similarities or differences between the mammalian MCR-1 and that of amphibians. An answer to this question may be forthcoming when the amphibian MCR-1 gene is cloned. In the meantime, we assume that the amphibian MCR-1 is present on the iridophore cell surface, but the distribution of this receptor on the surfaces of these differing pigment cells is unknown, and the question of whether these two chromatophores share identical proﬁles is unanswered. It is known through the use of various speciﬁc a-MSH antagonists that frog melanophores possess multiple MSH receptors, but such studies are in their infancy (Hruby et al., 1995). As such studies proceed, problems of species variation will be of concern. For example, an interesting study by Quillan et al. (1995) has revealed that D-Trp-Arg-Leu-NH2 is a potent a-MSH antagonist in Xenopus and that it is capable of causing pallor in adults when injected systemically. Moreover, it has the capacity to cause local lightening when applied top36

ically to such frogs. Xenopus is a rather particular amphibian; thus, the question arises of whether this MSH antagonist is generally active among amphibian species or is uniquely active in Xenopus.

The Role of Adrenergic Receptors While MSH seems to be the major factor regulating amphibian chromatophores, other hormones may also play a role in color change. Among these are catecholamines, such as epinephrine and norepinephrine, that apparently control the rapid color changes associated with “excitement pallor” (see Parker, 1948). Catecholamines are known to mediate certain effects through two types of receptors, a and b (Ahlquist, 1948), each of which controls responses that are antagonistic to the other. Accordingly, the response of a system to catecholamine stimulation depends on the presence or absence of a and b receptors, and it has been shown that both receptors may be present on amphibian chromatophores. With the aid of the receptor concept, it becomes possible to elucidate some previously unexplained paradoxical effects. For example, while both epinephrine and norepinephrine lighten the skins of R. pipiens by overriding the MSH effects (Wright and Lerner, 1960), catecholamines darken the skin of both Xenopus (Burgers et al., 1963) and Scaphiopus (Goldman and Hadley, 1969). Apparently, a receptors are present in R. pipiens (Lerner et al., 1954; Novales and Novales, 1965), accounting for the paling reaction, whereas in Xenopus (Graham, 1961; Novales and Davis, 1969) and Scaphiopus (Goldman and Hadley, 1969), b receptors predominate allowing darkening to occur in the presence of catecholamines. Although later advances in the pharmacology of a and b receptors have elaborated subclasses of these receptors, little has been done with respect to their deﬁnition in amphibians and reptiles. In contrast, the adrenoreceptors of ﬁshes have been better characterized in this regard (Fujii, 1993a). The color changes that occur in response to receptor stimulation in R. pipiens are attributable to changes in both iridophores and melanophores (Hadley and Bagnara, 1969). Moreover, the presence or absence of a and b receptors on iridophores and melanophores seems to complement one another. This is shown in the variation of response to catecholamine stimulation by different sibling species of R. pipiens (Hadley and Goldman, 1970). Norepinephrine lightens the MSH-darkened skins of R. pipiens of northern origin, but further darkens the skins of frogs of Mexican origin, probably Rana forreri. Lightening of the northern species is based not only on the presence of a receptors on the melanophores leading to an aggregation of pigment in these cells, but it appears that a receptors on the iridophore lead to dispersion of the reﬂecting platelets. In skins from southern frogs (R. forreri), b receptors predominate on both melanophores and iridophores so that catecholamine stimulation of skins in which MSH has already stimulated melanophores and iridophores leads to further melanosome dispersion in melanophores and greater reﬂecting platelet aggregation in iridophores. The simultaneous activation of the same receptor

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

type on both melanophores and iridophores does not always occur. For example, the catecholamine darkening of Scaphiopus skins mentioned above entails the activation of b receptors on only melanophores; although melanosome dispersion takes place in melanophores, iridophores remain unchanged (Goldman and Hadley, 1969). It seems reasonable that, as more studies are made relative to the application of the adrenergic receptor concept to pigment cell biology, it will be revealed that these receptors comprise an important part of the chromatophore system that can be used by a variety of agents known to affect chromatophores. An obvious question in this respect concerns the relationship between adrenergic receptors and cyclic AMP; does stimulation of adrenergic receptors lead to an alteration in the level of cyclic AMP within the chromatophore? It is tempting to speculate on this point in view of the report by Turtle and Kipnis (1967) that stimulation of b adrenergic receptors in tissues leads to an increase in tissue levels of cyclic AMP, whereas a adrenergic stimulation leads to a decrease in this substance. For a review of this topic, especially with respect to amphibians and reptiles, consult Bagnara and Hadley (1973) and Hadley and Bagnara (1975). An even richer literature has accumulated with respect to ﬁshes and has been cogently discussed by Fujii (1993a).

Molecular Mechanisms for Intracellular Translocation of Pigment Granules A variety of techniques and procedures have been utilized in order to comprehend the mechanism for intracellular transport of pigmentary organelles. In pharmaceutical studies of such motile responses, small pieces of skin or scales containing chromatophores were often used after removal of the epidermis by treatment with collagenase and/or chelating agents. For cell manipulation and microinjection of drugs and antibodies, in vitro cultured chromatophores were obtained as emigrants from skin explants or from tissues dissociated with collagenase and trypsin with subsequent centrifugal puriﬁcation in Percoll or Ficoll and immortalized as cell lines (Akiyama and Matsumoto, 1983; Akiyama et al., 1981, 1987; Aspengren et al., 2003; Daniolos et al., 1990; Fujii 1993a, b; Matsumoto et al., 1978, 1984; Negishi and Obika, 1980; Rogers et al., 1997, 1998). Electron microscopy discloses that ﬁsh and amphibian chromatophores contain an abundance of microtubules and actin ﬁlaments in a deﬁnite arrangement, together with relatively few intermediate ﬁlaments, the density and pattern of which are variable among species. When these cells are exposed to microtubule-disassembling agents such as colchicine, lumicolchicine, vinblastine, hexylene glycol, and nocodazole, as utilized in earlier studies, pigment aggregation is totally blocked or severely disrupted, mostly irreversibly and concomitantly with a disarrangement of their cytoplasmic framework (Obika, 1986; Obika and Negishi, 1985; Porter, 1973; Schliwa, 1984). When these cells are treated with drugs affecting actin ﬁlaments, viz. cytochalasins, DNase I, phalloidin, the results

are mostly confusing: in Xenopus and Rana melanophores treated with cytochalasin, dispersion is inhibited (Malawista, 1971; McGuire and Moellmann, 1972), whereas in ﬁsh melanophores exposed to cytochalasin B, pigment migration is not affected (Visconti and Castrucci, 1985). In goldﬁsh xanthophores, cytochalasin B inhibited heavy meromysin (HMM)-binding dispersion of carotenoid droplets (Lo et al., 1980; Obika et al., 1978). In swordtail erythrophores, microinjection of the anti-actin antibody clearly interferes with both aggregation and dispersion (Fig. 2.38) (Akiyama and Matsumoto, 1983). From these ﬁndings, it becomes clear that these two cytoskeletons are implicated in pigment translocation, more decisively for microtubules and with some diversiﬁcation for actin ﬁlaments. A unique aspect of this motility is bidirectionality with repeated centripetal or centrifugal migrations of the organelles occurring synchronously in whole dendrites of the cells. The question arises as to how these directed movements and their reversal occur. Inasmuch as microtubules have a polarity with minus and plus ends in their structure (Euteneuer and McIntosh, 1981; McNiven and Porter, 1986; McNiven et al., 1984), and inasmuch as they exist under a parallel alignment along the migratory tracks, as was indicated from pioneering studies (Bikle et al., 1966; Porter, 1973), these cytoskeletons are considered to be key instruments in this motility. As to motors driving pigment granules along microtubules, it was shown that cytoplasmic dyneins are inseparably implicated in aggregation as a minus end-directed motion whereas kinesins, mechanochemical ATPases, are involved in a plus end-directed dispersion (Asai and Lee, 1995; Haimo, 1996; Horikawa, 1998; Horikawa et al., 1998; Karki and Hozbaur, 1999; Rodionov et al., 1991; Thaler and Haino, 1996). The involvement of these motor proteins is primarily suggested by the requirement of this motility for a nucleotide triphosphatelike adenosine triphosphate (ATP) or guanosine triphosphate (GTP). This seems to be the case because the application of vanadate, which inhibits ATPase activity of dyneins, disrupts pigment aggregation in ﬁsh pigment cells (Beckerle and Porter, 1982; Clark and Rosenbaum, 1982; Luby and Porter, 1980; Negishi et al., 1985; Rozdzial and Haimo, 1986). Later, it was shown that cytoplasmic dyneins are colocalized with melanosomes along microtubules in both ﬁsh and amphibian melanophores (Nilsson et al., 1996; Rogers et al., 1997). Further decisive evidence supporting cytoplasmic dynein as the motor resides in the fact that puriﬁed melanosomes of Xenopus are capable of moving along microtubules towards the minus end in vitro (Rogers et al., 1997), and that microinjection of an anti-dynein antibody blocks pigment aggregation in ﬁsh melanophores (Nilsson and Wallin, 1997). In addition to these ﬁndings, it was also shown that microinjection of an antibody raised against kinesins (domain) into frog melanophores blocks pigment dispersion without affecting aggregation (Rogers et al., 1998). This suggests the possible participation of different motors in two phases of dispersion and aggregation. Biochemical analyses of puriﬁed 37

CHAPTER 2

A

D

B

E

C

F

melanosomes from Xenopus melanophores disclosed that kinesins are tightly associated with these organelles, moving them in vitro in the absence of cytosolic factors (Rogers et al., 1997, 1998). As kinesin II is a heterotrimer composed of two motor subunits of 85 and 95 kDa and a nonmotor of 115 kDa, mutations in these subunits should disrupt movement along microtubules. When a dominant-negative mutant of the 95 kDa motor subunit, which lacks the ATP-binding region of the motor domain, is produced, this headless kinesin II is unable to move organelles along microtubules (Tuma et al., 1998). In actin-based motility, myosin is considered to be the most reasonable candidate motor protein. As early as 1973, it was shown with mammalian melanocytes that myosin V, a nonmuscle myosin, is involved in melanosome transport from the cell center to the cell periphery where these melanosomes are transferred to keratinocytes (Wolff, 1973). In the dilute mutant of mice bearing defects in hair pigmentation, it is known that melanosomes are distributed in the perinuclear region of the melanocytes. Inasmuch as the dilute gene encodes myosin V, it is considered that melanosome transport along actin ﬁlaments toward the cell periphery is disordered by these mutated motor proteins, disrupting melanosome transfer from melanocytes to hair keratinocytes (Wu et al., 1997, 1998). 38

Fig. 2.38. The blockade of pigment displacement in cultured swordtail erythrophores by microinjection of an antiactin antibody. (A) Before microinjection. Cells are in a dispersed state in standard culture medium. (B) The same as (A) after administration of 5 ¥ 10-4 M epinephrine. All pigment (pterinosomes) is aggregated to the cell center. (C) The same as (B) after a brief rinsing in phosphate-buffered saline and subsequent administration of 10-3 M theophylline. Pigment is redispersed. (D) The same as (C) immediately after microinjection of the anti-actin antibody into a cell present in the center of the ﬁeld of view. A liquid parafﬁn droplet injected following the antibody solution is seen inside the cell body (arrow). (E) The same as (D) after exposure to epinephrine after injection. Note that pigment aggregation is blocked only in the injected cell, whereas uninjected cells nearby display complete pigment aggregation. (F) The same as (E) after exposure to theophylline. Note that no change is seen in the state of pigment dispersion in an injected cell, whereas pigment in the others is redispersed. Scale bar = 50 mm (from Akiyama and Matsumoto, 1983).

In Xenopus, it was shown with the use of isolated melanosomes that myosin V, although coexisting with kinesin II and cytoplasmic dynein, is tightly associated with these organelles, and that such isolated melanosomes are able to move along actin ﬁlaments in vitro (Rogers and Gelfand, 1998; Rogers et al., 1997). The localization of myosin V on melanosomes was also revealed in ﬁsh (cod) melanophores by immunoelectron microscopy combined with on-grid labeling using an antibody raised against 200 kDa mouse myosin Va heavy chain (Skold et al., 2002). The results of these studies strongly support the view that myosin V is implicated in pigment migration along actin ﬁlaments. Judging from the distribution pattern of actin ﬁlaments, which are rich at the cell periphery, it appears that motility based on their presence is of short-range nature. Physiological color change is mostly fast in ﬁsh and slow in amphibians, although fairly variable among species. In melanophores of the ﬁsh Oryzias latipes, the migratory speed of melanosomes is estimated as 0.06~0.3 mm/s as mass movement and 1~2 mm/s at 22∞C in an ideal linear track without a marked difference in the rate between aggregation and dispersion (Obika, 1986). In swordtail erythrophores (Fig. 2.39), pterinosomes migrate on average at 1 mm/s in aggregation and 0.01 mm/s in dispersion at room temperature, over about

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES A

50 µm

Aggregation(%)

B 100

50

Voltage 0.01mV

0 0

20 0 20 40 Time (min)

60

Fig. 2.39. Pigmentary responses of a swordtail erythrophore in an isolated scale. (A) Tracing of one cycle of pigment aggregation (left row) and dispersion (right row). The red pterinosomes are aggregated by administration of 5 ¥ 10-4 M epinephrine and dispersed by 10-3 M theophylline after washing in physiologic salt solution. The frame indicates the dimension of a recording photo cell. Note that there is little change in cell shape after one cycle of pigment responses. (B) Photoelectric recording of pterinosome displacement made on a single cell using a Rose chamber at room temperature (about 25∞C). The effects of yellow carotenoid vesicles are blocked by the use of a ﬁlter with a peak at 470 nm (see Matsumoto et al., 1984 for further details on instrumentation).

a 15~35 mm track in the dendrites (Matsumoto et al., 1984). In myosin-based motion, the velocity is estimated as about 0.04 mm/s, which would be appropriate for melanosome migration in amphibian melanophores (Tuma and Gelfand, 1999). The time needed for the completion of aggregation or dispersion would be affected by the proportion of processes using microtubule- or actin ﬁlament-based motors as well as the efﬁciency of their linkage. A proposal for the relationship of these two motility systems is depicted in Figure 2.40. It suggests that a microtubule-based motor works in long-range movement, utilizing cytoplasmic dynein for aggregation and kinesins for dispersion, while an actin ﬁlament-based motor, myosin V, acts in short range (Radionov et al., 1998; Wu et al., 1998). In the latter, pigment granules at aggregation are trapped at the cell periphery to be brought to microtubules for transport by cytoplasmic dynein, whereas at dispersion, pigment granules are dispersed evenly throughout the cytoplasm after transport from the cell center through microtubule-based motors (Tuma and Gelfand, 1999). This model is referred to as the “dual ﬁlament model of transport” by Langford (1995). Because ﬁsh chromatophores, when their actin ﬁlaments are drug disrupted, exhibit hyperdispersion possibly through a strong, centrifugal driving force of unaffected microtubule-based motors, and as frog melanophores under similar treatment show uneven distribution of melanosomes, possibly due to the lack of a local, short-range drive by a myosin motor (Radionov et al., 1998), it appears that this model is suitable for the interpretation of available ﬁndings. In signaling pathways for this motility, as mentioned earlier, the control of intracellular levels of cAMP plays a crucial role. This naturally suggests the involvement of protein phosphatases and protein kinases such as cAMP-dependent protein kinase A (PKA) or calcium-dependent protein kinase C (PKC) in this cascade (Nery and Castrucci, 1997). It is known that phosphatase 2A, but not 1 and 2B, is required for pigment aggregation of Xenopus melanophores, whereas phosphatase 2B, calcineurin, is necessary in ﬁsh (Reilein et al., 1998; Thaler and Haimo, 1990). In melatonin-induced aggregation of Xenopus melanophores, it was shown that mitogen-activated protein kinase (MAPK) is activated to supplement movement operated by the cAMP/PKA pathway (Andersson et al., 2003). On the other hand, it was shown that both PKA and PKC are implicated in pigment dispersion of Xenopus melanophores, despite the manner of their action being different and the role of PKC being supplemental. Activation of these two protein kinases is essential for dispersion of ﬁsh melanophores (Daniolos et al., 1990; Graminski et al., 1993; Reilein et al., 1998; Sammak et al., 1992; Sugden and Rowe, 1992). From these ﬁndings, it is apparent that phosphorylation and dephosphorylation of a target subunit of motor proteins is associated with regulation of bidirectional pigment transport (Rozdzial and Haimo, 1986). Current studies on this subject are focused on identiﬁcation of target subunits or phosphorylation sites of the motor proteins associated with the direction of migration. Knowledge of the role of calcium 39

CHAPTER 2 Aggregation

Dispersion Cytoplasmic dynein Kinesin II Myosin V

Melanosome Microtubule Actin filament

in this signaling cascade is still fairly controversial, depending upon the species used.

Cellular Associations in Color Change Alterations in the state of dispersion or aggregation of any single chromatophore type can often lead to profound color changes in an animal. However, color change often results from the integrated responses of the various pigment cells that exist together in well-organized associations. The association of one chromatophore type with another does not necessarily mean that either cell must undergo a physiological color change. Often, the association of two pigment cells is passive and serves to emphasize a permanent color pattern. As an example, the red spots on the adult dorsal surface of the newt, Notophthalmus viridescens, are based upon a precise superimposition of an erythrophore layer upon an iridophore layer (J. T. Bagnara, unpublished; Forbes et al., 1973). The red coloration of the spots is enhanced by reﬂection of light from the iridophores beneath. A similar situation applies to the bright spots of other species such as the spotted salamander, Ambystoma maculatum. Here, the yellow spots are based upon the exact superimposition of xanthophores upon iridophores in an otherwise homogeneous background of dermal melanophores (J. T. Bagnara, unpublished). There are many similar examples of chromatophore associations; however, the most important are the dermal chromatophore unit (Bagnara et al., 1968) and the epidermal melanin unit (Fitzpatrick and Breathnach, 1963). Neither of these units represents a precise structure in the anatomical and functional sense, but rather are concepts based upon the location of various chromatophores in the dermis and epidermis.

The Dermal Chromatophore Unit The primary function of dermal chromatophores is the pro40

Fig. 2.40. A model illustrating the implication of three motor proteins in melanosome translocation of ﬁsh and amphibian melanophores (adapted from Gross et al., 2002; Tuma and Gelfand, 1999; Wu et al., 1998).

duction of physiological color changes through the rapid intracellular mobilization of pigment-containing organelles. Among the adults of many amphibians, color changes are brought about by coordinated responses of the three basic chromatophore types, which are so situated that they comprise an integral, functional unit that has been designated the dermal chromatophore unit (Bagnara et al., 1968) (Figs 2.13, 2.18, and 2.41). More recently, a comparable unit has been described for ﬁshes (Fujii et al., 1989). Uppermost in the unit, just below the basement membrane, is a layer of xanthophores and immediately beneath this layer of yellow pigment is found a layer of iridophores. In frogs, the iridophore layer that forms the reﬂecting component of the unit is composed of a layer of melanophores that have dendrites extending upward. In frogs, these dendrites terminate in ﬁngerlike processes on the surface of the iridophore, just beneath the xanthophore layer. During adaptation to dark-colored backgrounds, melanosomes ﬁll these processes, thus obscuring the reﬂecting surface of the iridophore and leading to consequent darkening of the animal (Fig. 2.9). When the animal lightens, melanosomes move from the terminal processes and occupy a perinuclear position. As a result, their dermal melanophores are almost completely obscured by overlying xanthophore and iridophore layers and the animal appears light in color. The pigmentary role of the xanthophore layer relates to the establishment of the green color of many forms (Figs 2.23 and 2.42). In such animals, light waves leaving the iridophore surface appear blue because of Tyndall scattering and, as the light waves pass through overlying yellow pigment cells, the shortest wavelengths are absorbed so that ﬁnally the animal appears green (Fig. 2.17). The importance of the xanthophore pigments in imparting green coloration is shown not only by the blue coloration of green frogs from which yellow pigments have been leached (Fig. 2.9), but by the existence of blue mutants of frogs or

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

BL

X

PT

CV F

I RP

Fig. 2.42. Wholemount in Karo syrup of dorsal skin of a green tree frog, Hyla cinerea, photographed with epiillumination. Because of the presence of xanthophore pigments, green color is manifested. When yellow pigments are leached away, the preparation appears blue as in Fig. 2.9. The numerous black spots are openings of skin glands (see also Plate 2.12, pp. 494–495).

as larvae but not as adults when they have acquired the ability to change color rapidly. The loss of epidermal melanophores as metamorphosis approaches is paralleled by the differentiation of dermal chromatophore units.

M C MS

Fig. 2.41. Transverse section of Hyla cinerea skin from the dorsal surface showing dermal chromatophore unit in white backgroundadapted state. M, melanophore; I, iridophore; X, xanthophore. Melanosomes (MS) are uniformly distributed in processes extending around the sides of the iridophore, but ﬁngerlike endings (F) over iridophores are empty. BL, basal lamella; RP, reﬂecting platelets; CV, carotenoid vesicles; C, collagenous masses; PT, pterinosomes. Magniﬁcation ¥ 9700 (from Bagnara et al., 1968).

snakes (Fig. 2.22). In such mutants, the pigment content of the overlying xanthophores or erythrophores is almost completely depleted (Bagnara et al., 1978b). The effectiveness of dermal chromatophores and the dermal chromatophore unit as elements of physiological color change is very much affected by the presence of epidermal melanophores and epidermal melanin units. Because the latter lead to the deposition of melanin in overlying epidermal cells, physiologic color changes in the dermis beneath are obscured, and thus an inverse correlation exists between the degree of development of the two systems. In tree frogs, which undergo rapid and profound color changes, a well-developed system of dermal chromatophore units is present and epidermal melanin units are lacking. These frogs have epidermal melanophores

Subcellular Associations Over the past 25 years, it has been suggested on several occasions that the various kinds of pigment-containing organelles of dermal chromatophores are closely related to one another from the point of view of origin (for full details, see Bagnara, 1972; Bagnara and Ferris, 1971; Bagnara et al., 1979; Taylor and Bagnara, 1972). Essentially, it is believed that speciﬁc pigment-containing organelles, melanosomes, reﬂecting platelets, and pterinosomes may be derived from a common equipotential primordial organelle that may form, depending on speciﬁc developmental cues, any of the deﬁnitive organellar types. The strongest evidence in support of this conclusion is based upon the existence of chromatophores of one type that contain pigmentary organelles of another type (Fig. 2.43). So many examples of this phenomenon exist among amphibians that new observations are often not reported. It was ﬁrst noted that dermal melanophores of the canyon tree frog, H. arenicolor, sometimes contain a few reﬂecting platelets intermingled between melanosomes (Bagnara, 1972; Bagnara and Ferris, 1971; Taylor, 1971). Another example is represented in the skin from the dorsal surface of the red-backed salamander, Plethodon cinereus (Bagnara and Taylor, 1970). Erythrophores of the red form of this species contain pterinosomes for the most part but, in addition, a few melanosomes are found. Similarly, in melanophores of the dark form of this species, all three of these organelles are found. Also observed in erythrophores of this species are electron-dense organelles that may be mosaic intergrades between the melanosome and the pterinosome. The ﬁrst example of a 41

CHAPTER 2

C

P

RP

Fig. 2.43. A mosaic dermal chromatophore of the leaf frog (P. dacnicolor). Melanosomes, reﬂecting platelets (RP), pterinosomes (P), and carotenoid vesicles (C) are all present in the same cell (from Bagnara, 1983).

deﬁnitive normal pigmentary organelle containing at least two different unrelated pigments within the same limiting membrane is the earlier mentioned melanosome of adult P. dacnicolor, which contains both eumelanin and a pteridine pigment (Fig. 2.10). This species has also been shown to produce mosaic pigment cells and mosaic organelles of several types (Bagnara et al., 1979). Mosaic pigment cells and mosaic organelles are not limited to amphibians but have also been observed in reptiles and ﬁshes (Matsumoto et al., 1980). The general phenomenon of chromatophore mosaicism speaks to the obvious close developmental relationships among these neural crest-derived cells that are so obviously different in phenotype. It may also provide an explanation for the remarkable existence of chromatophore transdifferentiation, whereby a fully differentiated chromatophore of one type may change its phenotype and convert to a chromatophore of another type (see Ide, 1986). It appears likely that, as chromatophores of more species are examined, more examples of such chromatophore polymorphism will be observed, which may contribute to a fuller understanding of the mechanisms of chromatophore differentiation.

Pigmentation Patterns The striking and varied pigmentation patterns of poikilotherms have long attracted interest, and an understanding of their biological and physical bases has been sought after for 42

Fig. 2.44. Ventral surface of the same froglet which had received a neural fold graft at the neurula stage. Note the clearly circumscribed area of the graft and its pattern of atypical small spots. The original graft included prospective dorsal skin which had been already determined at the time of transplantation (see also Plate 2.13, pp. 494–495).

many years. Despite these efforts, substantial deﬁnitive knowledge about the development and maintenance of pigmentation patterns is lacking, although it is obvious that the ultimate expression of speciﬁc patterns depends upon the distribution of speciﬁc pigment cells. Pigmentation patterns may be general, such as the frequently encountered one in which the dorsum is darkly pigmented while the ventrum is light colored (Fig. 2.44 and see Fig. 2.49). The cellular basis for this pattern usually involves a greater presence of melanophores in the dorsum than in the ventrum and, conversely, a maximal presence of iridophores or leukophores in the ventrum and a minimal expression of these cells in the dorsum. More speciﬁc patterns result from the localized expression or lack of expression of selected chromatophores in circumscribed areas that are shaped as spots, stripes, mottlings, etc. These points are illustrated in Fig. 2.44 and see also Figures 2.47–2.49.

The Role of Endogenous Factors of the Integument The concept that patterns of chromatoblast differentiation are

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

Fig. 2.45. Dorsal surface of a newly transformed froglet of Rana pipiens showing the typical large spot pattern (see also Plate 2.14, pp. 494–495).

established early, even at the level of the neural crest before migration starts, has been considered for many years (see Bagnara, 1987); however, much recent experimentation has supported the alternative view that the pattern resides in the embryonic skin and that uncommitted chromatoblasts are inﬂuenced by speciﬁc local factors that inﬂuence the differentiation of these chromatoblasts (Bagnara et al., 1979). The ﬁrst concerted effort to discover factors of local integumental origin that might regulate chromatoblast differentiation was made by Fukuzawa and Ide (1988), who discovered a putative inhibitor of melanoblast differentiation in ventral but not dorsal skin of young Xenopus froglets. They considered that this agent present in ventral skin inhibited melanization of melanoblasts and thus designated it a melanization-inhibiting factor (MIF). They considered the differential activity of MIF in dorsal and ventral skin to be responsible for dorsal/ventral patterns. Later, Fukuzawa and Bagnara (1989) demonstrated that MIF (ventral conditioned medium) could override the stimulatory effects of trophic factors such as MSH that would normally be present in the living organism. They further showed (Bagnara and Fukuzawa, 1990) that putative MIF had

Fig. 2.46. The dorsal surface of a half-grown adult which had a portion of dorsal skin rotated 180° at a mid-larval stage. Note the exactness of ﬁt between the separated areas, indicating that the spot pattern was determined precisely at the time of rotation, when no indication of the pattern was yet evident (see also Plate 2.15, pp. 494–495).

a stimulatory effect on iridophores, as would be expected if MIF were indeed responsible for dorsal/ventral pattern formation (see Figs 2.47–2.49). The MIF molecule has been partially characterized, and a monoclonal antibody against MIF has been obtained (Samaraweera et al., 1994). Use of this antibody as an immunohistochemical probe has revealed that MIF is speciﬁcally localized in ventral skin of leopard frogs (Fukuzawa et al., 1995). The disclosure of an MIF from lower vertebrates coincides with the discovery that the agouti locus of mammals codes for a speciﬁc protein that similarly inhibits melanization of both mammalian melanocytes and melanophores of Xenopus and may be responsible for the dorsal and ventral pigmentation pattern of mice (Miller et al., 1993; Vrieling et al., 1994). The possibility that the agouti protein is related to MIF is a distinct one in view of the fact that agouti protein is an antagonist of the MSH receptor (Lu et al., 1994) and MIF blocks the melanogenic effect of MSH on Xenopus neural crest cells (Fukuzawa and Bagnara, 1989). 43

CHAPTER 2

Fig. 2.47. Iridophores of Pachymedusa dacnicolor in primary culture in control medium. Each iridophore has few processes and little or no branching of attached cells (see also Plate 2.16, pp. 494–495).

There is evidence that MIF has a similar effect on the mammalian system as it has been found that MIF extracted from the ventral skin of the leopard frog, Rana forreri, has an inhibitory effect on the activity levels of tyrosinase and dopachrome tautomerase in B16/F10 and Cloudman S-91 melanoma cell lines (Lopez-Contreras et al., 1996). Moreover, MIF seems to block the stimulatory effects of a-MSH on these enzymes. As MIF does not block the melanogenic effects of theophylline on these melanoma cells, it appears that it acts proximal to the MSH receptor. A similar search for a melanization-stimulating factor (MSF) has revealed the presence of such a putative factor in ﬁshes (Zuasti et al., 1992, 1993), and this factor has been partially characterized from catﬁsh skin (Johnson et al., 1992). What may be a similar factor has been demonstrated in the dorsal skin of the leopard frog where it appears to be particularly manifested in the dark spots (Mangano et al., 1992). Consistent with the view that the integument itself is responsible for the establishment of speciﬁc patterns is the demonstration by Bagnara (1982) that, in the leopard frog, dorsal ectoderm at the open neural plate stage has already been deter44

Fig. 2.48. A similar culture of iridophores in a medium conditioned by exposure to ventral skin of a leopard frog and presumably containing MIF. Note the large size of the iridophore mass containing conﬂuent cells and possessing many branching processes. MIF is presumed to stimulate iridophore differentiation (see also Plate 2.17, pp. 494–495).

mined as dorsal and can program immigrating chromatoblasts to differentiate in accordance with a dorsal spot pattern of a general type. He also showed that, at some point in early larval life, a highly speciﬁed ﬁnal spot pattern is put in place, long before it is expressed during metamorphic climax. Very likely, this speciﬁcation (or determination) takes place at the onset of the feeding stage (Naughten, 1971). It is attractive to consider the possibility that, at this early stage, factors such as MIF and MSF are expressed at a sufﬁcient level to affect chromatophore differentiation and thus to dictate the highly speciﬁed pigmentation pattern (see Figs 2.44–2.49 for details). The putative MIF and MSF molecules are presumably produced by cellular elements of the environment in which the pigment cells are found or pass through. In contrast, it is possible that patterns of migration and expression are inﬂuenced by molecules produced by the pigment cells themselves. In a study to investigate this possibility, Fukuzawa and Obika

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

(1995) considered that cell adhesion molecules (CAMs) expressed by speciﬁc pigment cell types might be important in poikilotherms. They discovered that, in both the medaka (Oryzias latipes) and Xenopus, N-CAM and N-cadherin are expressed speciﬁcally in xanthophores, but not in melanophores and iridophores. At present, it is too early to assess the meaning of these ﬁndings in the total expression of pigmentation patterns, but continuing studies hold promise of augmenting our knowledge in this area. It would be helpful if numerous pigmentary mutations were available in order to add a genetic component to our understanding of amphibian pigmentation patterns; however, only a few are adequately described, and even these are difﬁcult to utilize as development to sexual maturity in amphibians is relatively slow, and breeding colonies are restricted to only a few species, most notably the Mexican axolotl, Ambystoma mexicanum. Three pigmentary genes in this species, wild type (D), white (d), and albino (a), have been utilized in conjunction with microsurgical manipulations (chimera formation, reciprocal neural crest grafts, and grafts of gonadal primordia) to provide insights into an understanding of pigment pattern formation (Fig. 2.50). Houillon and Bagnara (1996) noted that “in chimeras between white and albino embryos, melanoblasts from the white half crossed the graft interface to differentiate in albino skin. Neural crest grafts from white embryos to albinos provided melanophores of white origin that were capable of differentiation in albino skin. Grafts of gonadal primordia from albino to white embryos provided albino germ cells that formed unpigmented ovocytes together with dark ovocytes: white ovocytes from the albino grafted ovary, and dark ovocytes from the host ovary. The donor albino white ectoderm included in the graft was able to support the differentiation of melanophores, iridophores, and xanthophores that invaded the graft ectoderm from the neural crest of the white host. It was concluded that manifestation of the white or wild phenotypes may be related to the possible presence or absence of inhibiting or stimulating pigmentary factors in the skin.” It seems possible that these respective inhibiting or stimulating factors are the circumscribed agents MIF and MSF that have been discussed.

Color Pattern Formation in Zebraﬁsh Integumental color patterns of ﬁsh are amazingly diverse even among mutants or variants of a single species (Kirschbaum, 1975; Yamamoto, 1975). It is no wonder then that biologists and ﬁsh fanciers alike are attracted to them: the former to elucidate the mechanisms responsible for the expression of particular patterns and the latter to produce new and interesting color forms or patterns. With the advent of the present age of molecular genetics, the zebraﬁsh, Danio rerio, and related species of this genus have become widely utilized as model organisms in studies on pattern formation. These species offer a wide choice of pigmentary mutants that are readily detectable through visible characteristics that are advantageous for the screening of mutations (Haffter et al., 1996). Integumental pigmentary patterns of adult zebraﬁsh are

Fig. 2.49. The margin of a graft of ventral larval skin of R. pipiens to the dorsal surface. Note that, after metamorphosis, the typical adult skin was expressed and thus a precise margin is manifested between the heavily iridophore-laden ventral skin and the dorsal surface. The former is considered to contain a melanization-inhibiting factor (MIF) that inhibits melanization and stimulates iridophore expression (see also Plate 2.18, pp. 494–495).

characterized primarily by melanophores that form either narrow, horizontal stripes on the body, as seen in the wild type, or dispersed discrete spots as in the panther or leopard mutants (McClure, 1999; Quigley and Parichy, 2002; Rawls et al., 2001) (Fig. 2.51). In other related species of this genus, such as Danio malabaricus, melanophores are also involved in the development of complicated reticulations over the ﬂank (McClure, 1999). All these color patterns of adults are formed after metamorphosis when the basic body structure switches from the larval to the adult form. In contrast to the diverse pigmentation patterns of wild-type or mutant adults of this genus, larvae display an essentially similar pigmentation pattern (McClure, 1999). Pigmentation of the larvae begins with the expression of melanophores in a few loose lines and is followed, with some delay, by the appearance of xanthophores distributed randomly. Based upon the onset of their appearance and cell size and density, 45

CHAPTER 2

A

B

A

B (a)

(b)

(c) C

(d)

(e)

Fig. 2.50. A. Two-year-old adult albino white chimeras. On the left chimera, note the clear line of demarcation separating the white (d/d) anterior half from the albino (a/a) posterior half. Some melanoblasts from the white half have migrated a short distance across the junction to differentiate and provide the black patches seen in the albino half. The same explanation applies to the chimera on the right except that, in this individual, white is posterior and albino anterior. Female on left, male on right (from Houillon and Bagnara, 1996). B. Two aD/aD albinos 2 years after having received neural crest grafts. Note the presence of melanophore patches distributed randomly in the dorsal area corresponding with the level of the neural crest graft. Female on left, male on right (from Houillon and Bagnara, 1996). C. A young adult female white Ad/Ad host 9 months after having received a gonadal primordium graft from an albino aD/aD donor. Note that the grafted albino integument is clearly delineated from that of the white host by its heavy pigmentation identical to that seen in wild-type individuals; both melanophores and iridophores are clearly visible. This female oviposited a majority of pigmented eggs corresponding with her dd genotype and a signiﬁcant number of white eggs derived from albino (a/a) germ cells contained in the graft (from Houillon and Bagnara, 1996) (see also Plate 2.19, pp. 494–495).

it is considered that there are two types of melanophores in these ﬁsh, larval and adult types. The occurrence of larval- and adult-type melanophores during ontogeny is well known for other teleost species (Matsumoto et al., 1960; Matsumoto, 1965b). In wild-type zebraﬁsh, adults are marked by several horizontal and regularly spaced melanophore stripes over the 46

(f)

(g)

Fig. 2.51. Schematic drawings of pigment (melanophore) patterns of zebraﬁsh, Danio rerio, and its related species. (A) A larva of the wild-type zebraﬁsh. (B) Adults: a, wild-type zebraﬁsh; b, panther (fms) mutant; c, sparse (kit) mutant; d, rose (endothelin) mutant; e, nacre (mitf) mutant; f, D. kerri; g, D. malabaricus (adapted from McClure, 1999; Quigley and Parichy, 2002; Rawls et al., 2001).

ﬂank. This pattern is known to form on the basis of two events: (1) intensive proliferation of adult-type melanophores at or after metamorphosis; and (2) expulsion of these melanophores by pre-existing xanthophores (Johnson et al., 1995; McClure, 1999). On the basis of the careful tracing of pigment cell development, McClure indicates clearly that adult-type melanophores appearing after metamorphosis are forced to coalesce, through contact with xanthophores, into lines along loosely aligned larval melanophore stripes, and that two populations of melanophores and xanthophores are gradually seg-

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

regated into two regions, narrow melanophore stripes with distinct margins and interspaces of xanthophores that are free of melanophores. The capacity of xanthophores to expel melanophores is clearly substantiated from observations of pigment pattern formation in the panther (fms), which lacks xanthophores in larval forms and in which stripes fail to form at metamorphosis. Instead, numerous tiny melanophorecontaining spots form over the adult body (Haffter et al., 1996; Johnson et al., 1995; McClure, 1999). It was also shown that the width of the black stripes, when xanthophores are present, depends upon the growth rate of adult-type melanophores appearing after metamorphosis and leading to the formation of wide stripes or large reticulations as a marked increase in their numbers occurs (Johnson et al., 1995; McClure, 1999; Quigley and Parichy, 2002). All these ﬁndings indicate that interactions between different pigment cell types, melanophores and xanthophores, and a genetically deﬁned proliferation rate of xanthophores play crucial roles in pigment pattern formation in these ﬁsh. In some related species such as Danio albolineatus and Danio sp. cf. aequipinnatus, erythrophores have been shown to play a role in pattern formation similar to that of xanthophores. An apparent “hostile” relationship between adult-type melanophores and xanthophores is considered to function in maintaining stripes in a deﬁnitive pattern (McClure, 1999). When melanophore migration along the dorso-ventral axis is disrupted locally in the course of stripe formation, the regularity of stripe alignment is lost, resulting in irregular-shaped reticulations or patches as seen in Danio kerri or D. malabaricus (McClure, 1999). Additional factors leading to such distorted stripes may include the development of neuromasts in the migratory pathways of melanophores and an uneven distribution of extracellular matrix. As to the apparent hostile relationship between adult-type melanophores and xanthophores, the possibility is suggested that it derives from their differing dependency on fms and kit in the course of development from common precursors, as the former requires either kit or fms, depending upon lineage, while the latter deﬁnitely requires fms (Mellgren and Johnson, 2002; Parichy and Turner, 2003; Rawls and Johnson, 2000). With respect to cell adhesion molecules of chromatophores, it was shown in the ﬁsh medaka (Oryzias latipes) and the amphibian, Xenopus laevis, that N-CAM and N-cadherin are speciﬁcally expressed in xanthophores, but not in melanophores and iridophores (Fukuzawa and Obika, 1995). If the same holds true for zebraﬁsh or its closely related species, such differences between melanophores and xanthophores may relate to a hostile relationship in their respective behaviors. It is interesting that this concept of chromatophore hostility, so well documented for zebraﬁsh in modern studies, is a reafﬁrmation of the views of Twitty and Niu (1954), voiced more than 50 years ago from work on newts. Recent studies on zebraﬁsh mutants with the use of the techniques of molecular genetics have disclosed that a variety of genes, most of which are orthologs of those in mammals, func-

tion in pigment cell development of this species and ultimately determine their pigmentation patterns (Haffter et al., 1996; Rawls et al., 2001). With respect to genes associated with melanophore development, the following mutations are known to be well-established examples (Fig. 2.51). In homozygous nacre (mitf ) mutants, no melanophores develop during embryonic, larval, and adult stages, indicating an absence of larval- and adult-type melanophores. In homozygous panther (fms) and rose (ednrb1) mutants, the proliferation of adulttype melanophores associated with body stripe formation is reduced, as mentioned previously, although fms and enrb1 are ortholog genes encoding, respectively, a type III receptor tyrosine kinase and a G-protein-coupled endothelin receptor B in mammals. In homozygous sparse (kit) mutants, the development of adult-type melanophores to be incorporated into body stripes and dorsal scales is disrupted. In double sparse (kit) and rose (ednrb1) mutants, two types of larval- and adulttype melanophores fail to develop, while a certain number of melanophores appear in the caudal and anal ﬁns. In view of the development of caudal and anal ﬁn melanophores in sparse and rose double mutants, even in the absence of larvaland adult-type melanophores in the ﬂank, and in light of the development of melanophores having different requirements for kit in regenerating ﬁns, it seems that four different classes or lineages exist in zebraﬁsh (Rawls et al., 2001). Some of these genes associated with the fate of melanophores of this species are also implicated in the development of other types of chromatophores as exempliﬁed by fms for adult-type xanthophores and ednrb1 for adult-type iridophores (Rawls et al., 2001). In addition, a group of genes such as pfeffer (pfe) and salz (sal) causes reduction in xanthophore populations, whereas another group including rose (rse), shady (shd), and transparent (tra) does the same for iridophores (Haffter et al., 1996; Rawls et al., 2001). With respect to mutations affecting melanogenesis itself in adults, a number of genes including albino (alb), mustard (mrd), and sandy (sdy) are reported to be incapable of supporting the production of melanin, and a group of brass (brs), fading vision (fdv), and golden (gol) express a phenotype of reduced melanin (Haffter et al., 1996). As studies of the molecular genetics of zebraﬁsh pigmentation are largely facilitated by those on mammals, available information on pigment genes of this species is rather restricted to melanophores. The information so far derived indicates that the functions of pigmentation-associated genes in zebraﬁsh are in essence comparable to those in mammals, except for the occurrence of a variety of melanophore lineages in this species. In view of the fact that Wnt-1 and Sox 10, essential for the development of mammalian neural crest, are expressed in developing melanophores of zebraﬁsh (Rawls et al., 2001), it is evident that a common basis is shared in the development of pigment cells of both groups. Knowledge about pigment cells of lower vertebrates certainly provides a useful key for understanding the mechanisms of pigment pattern formation and its related phenomena in the vertebrate kingdom. 47

CHAPTER 2

Hormonal Inﬂuences on the Development of Pigmentation Patterns Because of its importance in morphological color change, MSH would seem to be a likely candidate as a major role player in pigment pattern formation. As was pointed out earlier, MSH exerts a powerful proliferative effect on melanophores (Pehlemann, 1967a, b, 1972) and, as shown in Figure 2.30, apparently stimulates the differentiation of latent melanoblasts (Bagnara and Fernandez, 1993). Its stimulatory effects can be so potent on some frogs that the normally pale ventrum is darkened (Figs 2.27 and 2.28) (Fernandez and Bagnara, 1991, 1993). The morphological effect stemming from the lack of MSH must also be considered when evaluating the expression of pigmentation patterns. In this case, iridophores are maximally expressed in the absence of MSH. As was pointed out earlier, Xenopus larvae deprived of MSH from the earliest larval stages develop iridophores in the tail ﬁn (Fig. 2.26) where they normally never appear (Bagnara, 1957). One of the best examples of the inﬂuence of MSH on the expression of normal pigmentation patterns is that in wild isogenic populations of tiger salamanders, in which the environment markedly affects the amount of MSH secreted and thus the pigmentation pattern that is manifested (Fig. 2.33) (Fernandez and Collins, 1988). High-albedo environments result in low MSH levels and the high expression of iridophores that mask melanophore inﬂuences on pigmentation patterns. Low-albedo individuals secrete high levels of MSH and thus a pattern dominated by melanophores is expressed. As potent as these effects of MSH are on morphological color change, they nevertheless must not be construed as an indication that MSH is a primary determinant of pattern formation (Bagnara and Fernandez, 1993). In fact, a clearly formed pigmentation pattern of dorsal spots is manifested in leopard frogs hypophysectomized as embryos, but induced to metamorphose (Fig. 2.52). The role of MSH is not a causative one in the development of primary pigmentation patterns; rather, it is a modiﬁer through its actions on the differentiation of latent chromatoblasts or on proliferation. Similarly, thyroxine plays a permissive role in pigment pattern formation through its general action on the skin, and thus on pigment pattern expression, at metamorphic climax (Bagnara, 1982).

Pigmentation Anomalies Obviously, in nature many undiscovered pigmentary abnormalities exist. These range from albinism and other color mutations to serious pathological conditions such as pigment cell malignancies. With the tremendous growth in the popularity of ﬁshes, amphibians, and reptiles as pets, a large industry has grown up in the breeding of these poikilotherms, and color variants have been selected for. Unfortunately, the professional research community has not taken advantage of these variants for research, in large measure because of the lack of ﬁnancial support for such seemingly esoteric studies. In contrast, pigment cell malignancies have been the focus of inves48

Fig. 2.52. A R. pipiens tadpole in metamorphic stasis due to an anomalous development of the hypophysis such that MSH is lacking and TSH is insufﬁcient to induce metamorphic climax. Note the obvious onset of the formation of a typical spot pattern even in the absence of MSH.

tigation for many years and, in fact, the establishment of pigment cell biology as a recognized discipline (Bagnara, 1991) stems from the discovery of the Xiphophorine GordonKosswig Melanoma System. This subject has been reviewed frequently in recent years by Professor Fritz Anders, and thus the reader is referred to one of his important works (Anders, 1991). A lesser known malignancy is erythrophoroma from goldﬁsh, an erythrophore neoplasm ﬁrst described by Prince Masahito and his collaborators (Ishikawa et al., 1978), who ultimately established several lines of neoplastic erythrophores (Matsumoto et al., 1980) (Fig. 2.53). Later, uncloned cell lines of iridophores and melanophores were derived from an iridophore–melanophore tumor found on a marine teleost, the nibe croaker (Matsumoto et al., 1981). This neoplasm (iridomelanophoroma) metastasized on both dorsal and ventral regions of the ﬁsh with the melanophore component more prevalent in the dorsum and the iridophore phenotype more prevalent in the ventrum. It is tempting to speculate that the two phenotypes appear where they do as a result of the actions of MSF and MIF respectively. A similar neoplasm has been described from the pine snake (Fig. 2.54) (Jacobson et al., 1989). It was referred to as an iridosarcoma, but it appears very much like the irido-melanophoroma and has the capacity to transform from a largely iridophore type to one that is more phenotypically melanophore-like. Although the malignancy is primarily dermal, neoplastic cells seem to traverse the basement membrane with ease to invade the epidermis (Fig. 2.55). Lower vertebrate pigment cell neoplasms (chromatophoromas) have much to offer in basic research; however, little is being done with them at present. Very likely, there are numerous such malignancies that appear both in nature and in the colonies of ﬁsh, frog, and reptile breeders.

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

Fig. 2.53. Morphological properties of spontaneous cutaneous erythrophoromas and irido-melanophoromas occurring in teleost ﬁsh. Left column, erythrophoroma formed on the dorsal surface of an adult goldﬁsh. Note that the tumor contains several growing nodules (middle). Component cells of the tumor, even in the central portion, are pigmented red to orange as are normal erythrophores present in the skin. Right column, the proﬁles of cells from the GEM 81 cell line of goldﬁsh erythrophoromas (top), and nibe croaker irido-melanophoroma (middle and bottom) exhibiting either melano(phoro)ma-like pigmentation (middle) or iridophoroma-like cells (bottom) depending upon the site of the lesion. Scale bars = 100 mm (see also Plate 2.20, pp. 494–495).

Unfortunately, these usually escape the attention of pigment cell investigators; however, the case of the aforementioned goldﬁsh erythrophoroma is an exception, for its discovery followed by appropriate investigation has led to valuable contributions to our knowledge of pigment cells. Interesting aspects of goldﬁsh erythrophoroma cells are

their instability or multiplicity of phenotypic expression. The vast majority of erythrophoromas in situ are recognized by their yellow or orange pigmentation as a tumor of erythrophores, although some may contain a few black lumps composed of numerous melanophore-like cells (Ishikawa et al. 1978; Matsumoto et al., 1980). When these erythrophoroma 49

CHAPTER 2

cells are cultured in a standard culture medium in vitro as a permanent cell line derived from a sporadic tumor of orangecolored goldﬁsh, they remain essentially unpigmented, although they may become pigmented following the addition of ﬁsh serum to the culture medium (Matsumoto et al., 1980; 1983). When these cells are subjected to factors that induce differentiation, viz. the use of carp serum in combination with

Fig. 2.54. An adult northern pine snake (Piterophia) with irregularly thickened and pitted black and white ventral scales. A large subcutaneous iridosarcoma nodule bulges to overlying skin (arrow) (from Jacobson et al., 1989).

dimethyl sulfoxide (DMSO) or others, they begin to form, mostly on a clonal basis, a variety of products such as melanin, reﬂecting substance, teeth- or bone-like structures, and lenslike bodies (Matsumoto et al., 1981, 1983, 1989) (Fig. 2.56). Cells expressing an abundance of melanin are essentially similar to melanophores with respect to both their dendritic appearance and their immense deposition of melanosomes and, upon clonal subcultivation, develop a responsiveness to aggregating or dispersing agents such as epinephrine or cAMP respectively (Matsumoto et al., 1982, 1989). Cells that contain a reﬂecting substance appear similar in proﬁle to iridophores, as evidenced by the presence of bizarre-shaped platelets (Matsumoto et al., 1981, 1989). Within cell mounds that develop in long-sustained cultures of these cells are lentoid bodies that form occasionally and in which crystallins are detectable by immunochemical assay (Akiyama et al., 1986). In other cases, teeth-like structures are recognized within such cell mounds, suggesting their differentiation toward odontoblasts or the like (Matsumoto et al., 1983). In some cultures, neuron-like cells appear, extending thin, long dendrites that lack pigment deposition (Matsumoto et al., 1983). As most of the cell characteristics that are thus expressed are of neural crest origin (Weston, 1970), it is considered that immortalized erythrophoroma cells are neural crest stem cells in nature. It seems likely that previously described differentiations such as schwannomas or neuroﬁbromas, etc. deriving from goldﬁsh tumors (Duncan and Harkin, 1969) are manifestations of the multiplicity of possible phenotypic expression that these tumor cells are capable of. The capacity for multiple differentiation exhibited by both goldﬁsh erythrophoroma cells and nibe iridomelanophoroma cells offers a wide potential for their use in studies of pigment cell differentiation.

Fig. 2.55. Dermal epidermal junction from an abnormal black scale from the affected region shown in the previous ﬁgure. Note the many abnormal iridophores (left) and a single iridophore in the corresponding epidermis just above the basement membrane (arrows). Magniﬁcation ¥ 3700 (modiﬁed from Jacobson et al., 1989).

50

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

Fig. 2.56. Multiple differentiation of goldﬁsh erythrophoroma cells in vitro. Top, clone of melanophore-like cells that are capable of exhibiting pigmentary responses such as melanosome displacement from a dispersed state (left) to an aggregated state (right). Aggregation is induced by the addition of epinephrine at 5 ¥ 10-4 M. Arrows indicate corresponding sites of the clone. Middle, clone of iridophore-like cells that exhibit brownish proﬁles under a transmitted light (left) and reﬂectance under reﬂecting illumination (right). Bottom, formation of tooth-like (left) structures (arrows) and bone-like (right) structures (arrows) within a cell mound. Differentiation is induced by the administration of dimethyl sulfoxide (DMSO) and ﬁsh serum (see Matsumoto et al., 1989 for details). Bars indicate 50 mm for top/middle and 25 mm for bottom (see also Plate 2.21, pp. 494–495).

Perspectives Much of the knowledge discussed herein has accumulated during the last 50 years and, given the low number of scien-

tists engaged in the area of pigment cell research on lower vertebrates, there is a wealth of untapped knowledge to be gained. In order properly to take advantage of the opportunities offered by these lower vertebrates, investigators engaged in research on mammalian pigmentary problems must realize 51

CHAPTER 2

that the pigmentary phenomena studied in lower vertebrates are not necessarily unique to these lower forms. Lower vertebrate pigmentary problems should not be studied merely because, by analogy, they offer model systems for relatively distant mammalian or human phenomena. It must be realized that, frequently, homologies exist among the pigmentary phenomena displayed by lower and higher vertebrates alike. After all, the pigment cells of all vertebrates are of neural crest origin, and they migrate in a similar manner to distal sites where they respond to similar factors present in either the immediate area they occupy or in the organism as a whole. An exceedingly important problem about vertebrate pigment cells that needs study is that of chromatophore organellogenesis. The few studies available have indicated that melanosomes, reﬂecting platelets, and pterinosomes are of endoplasmic reticular origin (Bagnara et al., 1979), but little is known about details. Unlike the situation with respect to melanosome formation, which has been much studied for mammals and can thus be extrapolated to lower vertebrates, there is no such possibility for iridophores and xanthophores. Perhaps the reverse will be true one day when it becomes appreciated that vestiges of xanthophores and iridophores are present in the iris of homeotherms, which may serve as a refuge for these chromatophores that have long been considered to be unique to poikilotherms (Oliphant et al., 1992). A related problem and one ultimately related to the question of chromatophore characterization and identiﬁcation concerns the mysterious pigmentary organelles that have been found in chromatophores of little-studied species such as the Centrollenid and Dendrobatid frogs mentioned earlier. A better understanding of these organelles can only be derived from studies on species of less recognized taxa. Probably, the existence of some of the unusual pigmentary organelles is related to speciﬁc ecological needs of the various taxa. Certainly, background color matching is an important antipredator behavior; however, it is possible that some of the unusual organelles contain pigments that help in other means of protection from predators. For example, Schwalm et al. (1977) have shown that some leaf-sitting frogs reﬂect infrared, and they suggest that this ability may confer advantages to the frogs with respect to both thermoregulation and infrared cryptic coloration. In this regard, some of the unusual organelles of poison arrow frogs, the Dendrobatids, may contain pigments that provide the bright warning colors typical of this group. Because of the rather favored status of ecological studies during present times, it is not unlikely that the role of chromatophores and their pigmentary organelles in topics such as thermoregulation, osmoregulation, predator avoidance, and reproduction stand a good chance for study. This is in contrast to the possibilities of studying chromatophores, merely for the sake of knowledge, for such investigations are too esoteric for these modern times. One of the intriguing aspects of chromatophore physiology that certainly needs investigation is that of photoreception. The suggestion that we made, almost 50 years ago, that

52

amphibian chromatophores may possess visual pigments (Bagnara, 1957), seems to have been vindicated by the demonstration that piscine chromatophores do, indeed, seem to possess visual pigments (Lythgoe and Thompson, 1984). However, this discovery seems to have escaped the attention of investigators of higher vertebrate pigmentation and of photobiology. Sooner or later, this knowledge will strike home, especially because of possible implications in the photobiology of tanning or in oncology. It would be fascinating, indeed, if it were found that some chromatophores of all types contain visual pigments, either those already characterized from the retina or entirely new types. Moreover, given the absence of the usual photoreceptive membranes typical of photoreceptors, it would be interesting to know whether the limiting membranes of the pigmentary organelles, themselves, serve as the surface repository for such pigments. Of possible relatedness to the question of direct light reception by chromatophores is the extracutaneous distribution of such cells. In one species or another, chromatophores occur in practically every internal organ, on blood vessels, body cavity linings, meninges, etc. It has been suggested that the presence of pigment cells and melanin in internal organs is related to the impact of solar radiation. It was beyond the scope of this chapter to delve into this subject, especially as much of the work is older and speculative; however, in this regard, two possible functions for these internal pigment cells stand out, thermoregulation and protection. As an example of the former, in a fascinating study by Guillete et al. (1983), it was discovered that one testis in several related species of spiny lizards is heavily melanized and is thus warmer. As a consequence, its spermatogenesis is accelerated over that of its partner. From the standpoint of protection, the presence of melanin in the kidney (Zuasti et al., 1989) and the liver (Sichel, 1988) has been viewed as being part of antioxidant systems based upon the reducing action of melanin. There is so little work being done in this area on lower vertebrates that it stands out as an important target for future investigation. Probably the area of research that holds the most promise for investigators of lower vertebrate pigmentation is that of pattern expression. In particular, the extrapolation of results stemming from mammalian molecular genetics to these lower forms has the most to offer. Earlier, it was commented upon that the putative MIF from amphibians is analogous to the agouti protein derived from the cloning of the gene from the mouse agouti locus. As can be seen from later chapters in this volume, several mammalian genes dealing with pigmentation have been characterized, and an array of mutations affecting mouse coat color have been described (see Jackson, 1994). Among the affected genes are those that affect melanocyte development and migration, melanocyte morphology, melanosomal structure and function, the enzymes of melanogenesis, and the MSH receptor. These characters are elements of pigment cell biology common to all vertebrates. Thus, it remains for the lower vertebrate counterparts of these mammalian genes to be found. Analyses of piscine and amphibian

COMPARATIVE ANATOMY AND PHYSIOLOGY OF PIGMENT CELLS IN NONMAMMALIAN TISSUES

tyrosinase genes and their products are well under way (Miura et al., 1995; Morrison et al., 1994; Peng et al., 1994; Takase et al., 1992; Yamamoto et al., 1992); however, genes dealing with pattern-speciﬁc products have not been extended to lower vertebrates. As mentioned earlier, attention to the nature of MSH receptors of lower vertebrates is just commencing. Such studies need to move forward with urgency given the important and varied roles of MSH in lower and higher vertebrates alike. The recent discovery by Valverde et al. (1995) that alterations in the MSH receptor markedly affect the pigmentary phenotype of man should provide special impetus to studies of MSH receptors and the genes that govern them in lower vertebrates. It is important to remember that the pigmentary systems of higher vertebrates are homologous to those of lower forms; thus, investigators involved in research at each of these levels should keep open minds and be able to apply knowledge obtained at one level to pigmentary problems of vertebrates at another level.

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The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

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The Science of Pigmentation

The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

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General Biology of Mammalian Pigmentation Walter C. Quevedo Jr. and Thomas J. Holstein

Part I: The Melanin Pigmentary System of Postnatal Mammals from Basic Design to Evolutionary Origins Part II: The Development of the Melanin Pigmentary System and the Molecular Genetic Systems that Regulate It

Summary 1 This chapter examines the structure and function of the mammalian pigmentary system in the light of its diversity, evolutionary origin, and contributions to the adaptiveness of mammals to natural environments. 2 A major focus is on the variation in skin color based on the interactions between melanocytes and keratinocytes in the pigmenting of the epidermis. 3 The inborn constitutive melanin pigmentation of human skin is contrasted with facultative melanin pigmentation or tanning in response to solar radiation. 4 As in the epidermis, melanocytes of hair follicles synthesize melanin and transfer it to keratinocytes (epidermal cells) that form the hair proper. 5 With respect to hair growth, follicular melanocytes that pigment each generation of hair appear to die at the end of each cycle and are replaced in the next one by melanocytes recruited from a stem cell population situated in the upper hair follicle. 6 Dermal melanocytes of humans are most frequently found at sites where hair is thin. “Mongolian spots” serve as a general model for the colors generated by dermal melanin. 7 The presence of dermal melanin is highly variable in mammals, suggesting that dermal melanocytes may have functions in addition to providing photoprotection. 8 Although mammals lack the capacity for rapid color changes by shifting pigment within melanocytes, the color interactions between static deposits of dermal melanin and surrounding blood vessels colored red by hemoglobin produce rapid ﬂashes of bright blue color in such primates as the mandrill (Mandrillus sphinx). 9 At least seven adaptive functions have been proposed for melanin in protecting humans from environmental hazards and for promoting effective metabolic functions of skin exposed to solar radiation. 10 The effectiveness of melanin in photoprotection is being debated. The big picture suggests that melanin is photoprotective to the extent that it is in harmony with its other functions.

11 The evolution of very light-skinned humans created the paradox of cutaneous melanin potentiating solar damage rather than providing photoprotection. 12 As an interface, the hair coat provides many signals to observers when used as communication or none at all when used as camouﬂage. The pigmentation of the hair coat is carefully crafted for these purposes. 13 Among mammals, youth is a time to play and learn. It is a time for protection from aggression from the adults that surround them. Coat colors signal to adults the privileged status of their conspeciﬁc juveniles. The coat colors of juveniles also elicit protection by their adults when predators threaten. In some cases, the coat colors of juveniles, rather than calling attention to them, facilitate concealment. 14 Radicalism and conservatism are evident in the human pigmentary system evolved from that of reptiles. Human skin retained certain features of the spatial organization in which melanocyte function is integrated into the functions of skin as a whole. Examples of this integration are melanin’s role in thermoregulation, the paracrine activity of melanocytes in the inﬂammatory response, and melanin’s neutralization of certain cytotoxic agents. 15 Many of the most basic features of the mammalian pigmentary system have been carried over from reptiles (melanocytes, melanosomes, tyrosinase, melanin, etc.). The major evolutionary change was the evolution of new regulatory mechanisms for orchestrating both genetically and epigenetically the patterns of pigment distribution in the skin and hair to facilitate the survival of mammals in an acutely visionoriented world. The details of the embryonic and postnatal settings in which the melanin pigmentary system is formed and integrated with local surrounding cells and distant glands that regulate it indicate the great importance of melanocytes in the rise of mammals to dominance in the natural world.

Historical Background The evolution of mammals from reptilian ancestors was characterized by the invention of a new system of protection, a 63

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coat of hair. This innovation required a revision of strategy for displaying skin color. Unlike their reptilian, amphibian, and ﬁsh ancestors, mammals could no longer proﬁt by using dermal chromatophores (xanthophores, melanophores, and iridophores) to produce stable or rapidly changing patterns of skin pigmentation, for their skin was no longer completely visible. In fact, the ability of chromatophores rapidly to disperse or concentrate pigments within their cytoplasm became a liability in mammals. In their case, the strategy for protective coloration had to be shifted from patterning of the skin with a variety of pigments to one of programming the pigmentation of hairs that grew from hair follicles. The pigment used for this purpose was melanin deposited within melanosomes, specialized organelles synthesized by melanocytes. Each hair had to be pigmented precisely from top to bottom to make its appropriate contribution to the pattern expressed by the dead hair coat that invested the skin. There was no place in mammals for disruptive reversals in the movement of melanosomes within melanocytes in response to endocrine or neural stimuli. Although such physiologic color changes were of great importance in lower vertebrates, they were nonadaptive in the hair follicles of mammals. In this location, unscheduled reversible shifts in the distribution of melanosomes within melanocytes would have interrupted the programmed synthesis and transfer of melanosomes from melanocytes into cells (keratinocytes) being incorporated into the developing hair. Any disruption in the program for pigmentation of small populations of individual hairs could produce a serious ﬂaw in the visual impact of the overall coat color pattern. Although melanocytes lost their capacity rapidly to disperse and concentrate their melanosomes in response to hormones, hormones did retain the ability to regulate pigmentation, hence hair color, by inﬂuencing the amount and type of melanin [eumelanin (black/brown) vs. pheomelanin (yellow/red)] synthesized by follicular melanocytes. Physiology deals with the functions of living organisms with particular attention to the mechanisms that regulate them at all levels of biological organization, from subcells to social groups and external environmental inﬂuences. In many ways, the mammalian pigmentary system is an ideal example for afﬁrming this interpretation of nature. Accordingly, this chapter emphasizes two approaches to exploring the melanin pigmentary system of mammals. Part I, starting from the top down, from complexity to “simplicity,” describes the big picture of the gross and microscopic organization of the melanin pigmentary system, its universal and variable features among mammals, its adaptive signiﬁcance, evolutionary origin, and the constraints placed on it by its inheritance from reptilian ancestors. Part II begins from the bottom up, from “simplicity” to complexity, in describing the shaping of the melanin pigmentary system during embryonic development. It stresses the roles played by events at the molecular level that regulate development, including the genes and their products, enzymes, hormones, and growth factors that ﬁne-tune the life, differentiation, and death of melanocytes. The interplay (inductions) of the epidermis and dermis prior to the devel64

opment of hair follicles unleashes the appropriate genetic mechanisms that specify and regulate the path that growth will take. Some features of the melanin pigmentary system are important in both mammalian evolution and the molecular biology of development. This chapter examines the links between them. In doing so, it crosses the boundary between physiological genetics and molecular genetics of the mammalian pigmentary system. In general, molecular genetics has focused on the central dogma of DNAÆmRNAÆprotein, whereas physiological genetics has emphasized the path from the gene product, a protein, the initial phenotype, to the derived phenotype, in this case the melanin pigmentary system that emerges during embryonic and postnatal development (Quevedo and Holstein, 1992). In an attempt to give an integrated picture of events at the micro level, some repetition of information about the macro level has been included to expand what was stated earlier in another context. At the end of the chapter, there is a list of other chapters in this book that expand on the topics covered here.

Part I: The Melanin Pigmentary System of Postnatal Mammals from Basic Design to Evolutionary Origins To begin with, the human melanocyte system will be used to illustrate the general principles of mammalian pigmentation. More is known about the melanin pigmentation of humans than for any other mammal, except for the laboratory mouse (Mus musculus). In general structure and function, the human pigmentary system shows little difference from that of other mammals. However, unlike other mammals, the marked reduction of hair that characterized human evolution once again placed the epidermis and not the hair coat as the primary interface between the organism and its environment. In addition, while hemoglobin imparts signiﬁcant color to human skin, its impact is not as great as in certain other mammals. Therefore, to complete the generalized picture of mammalian pigmentation based on Homo sapiens, (1) mice will be recruited for an analysis of the pigmentary system in skin sprouting dense hair and (2) the mandrill, a baboon-like primate, will be examined for it epitomizes the ability of the circulatory and pigmentary systems to cooperate in producing bright patches of red and blue in glabrous skin.

Current Concepts Human Skin Color In humans, skin color derives mainly from the interplay of two pigments, hemoglobin and melanin. Eumelanin in the epidermis typically imparts a brown to black hue, whereas eumelanin in the dermis may appear blue. However, the epidermis of orange–red-haired, fair-skinned, freckled individuals may

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

appear slightly orange–yellow, possibly owing to the presence of pheomelanin in higher proportions than eumelanin than is characteristic of individuals lacking red hair and freckles (Thody et al., 1991). However, recent ﬁndings by Hunt et al. (1995) cast doubt on this interpretation. Oxygenated hemoglobin in the superﬁcial dermal arterioles and capillaries appears pink/red and deoxygenated hemoglobin in the venules blue. Dietary carotenoids may contribute a slight yellow tint to the epidermis (Jimbow et al., 1999). For most humans alive today, melanin is the major source of skin color. Although melanocytes are found in hair follicles and occasionally in the dermis, worldwide ethnic variation in heritable (“constitutive”) skin color results mainly from differences in the amount of melanin within the epidermis. Constitutive skin color (CSC) is deﬁned as the level of melanization generated within the epidermis of an individual through the operation of cellular genetic programs in the absence of inﬂuences from ultraviolet light. In practical terms, it is taken to be the level of pigmentation in those parts of the human body that are rarely exposed to sunlight. Humans vary in CSC from dark black/brown to almost alabaster white in skin color. CSC can be enhanced by exposure to ultraviolet light (UVR). Facultative skin color (FSC) designates increases in melanin pigmentation of the skin above CSC and is induced by UVR and/or hormones. FSC is reversible in that the hyperpigmentation tends to decline toward CSC when solar and/or endocrine stimulation is discontinued (Quevedo et al., 1974).

The Histological Basis for CSC and FSC: Epidermal Melanocytes Human epidermis is conventionally described as a stratiﬁed squamous epithelium consisting of keratinocytes that are modiﬁed as they ascend from their origin in the basal layer and ultimately cornify and die as they near the outer surface where they are shed (Bloom and Fawcett, 1962) (Fig. 3.1A). However, the epidermis is a considerably more complex community of cells than this deﬁnition implies (Montagna and Parakkal, 1974; Odland, 1991). Although keratinocytes are the principal cells of the epidermis, there are also several types of nonkeratinocytes. Particularly prominent are the melanocytes found in the basal layer of the epidermis (Fig. 3.1A and B). Epidermal melanocytes typically synthesize melanosomes that become ellipsoidal in shape and more or less uniformly pigmented by melanin deposited on an internal matrix of aligned ﬁlaments (cf. Chapter 7). The melanin produced by epidermal melanocytes is most often described as eumelanin based on its tendency to appear brown/black in color. The mature melanosomes are moved from the cell body (perikaryon) of the melanocyte to its dendrites that are in contact with neighboring keratinocytes of the Malpighian layer (cf. Chapter 8). The melanosomes move from the dendrites into the keratinocytes. There are three major theories about how melanosomes are acquired by keratinocytes: (1) keratinocytes phagocytize bits of the melanosome-containing dendrites; (2) melanosomes are discharged by the dendrites into the intercellular space and subsequently phagocytized by

SC SS SG

D

A K

MC

MN

G G K

RER B

*

D

Fig. 3.1. Vertical section of skin from the buttocks of a heavily pigmented adult African American male. (A) Note the heavily pigmented keratinocytes of the epidermal basal layer. In some keratinocytes, the arrangement of melanosomes in the form of supranuclear caps is evident (arrowheads). A “clear cell” (arrow), presumably a melanocyte, is present in the basal layer of the epidermis (SC, stratum corneum; SG, stratum granulosum; SS, stratum spinosum; D, dermis). (B) Ultrastructure of a basal melanocyte ﬂanked by two keratinocytes. Melanosomes at various stages of development (arrows) are distributed throughout the cytoplasm of the melanocyte. Arrowheads indicate melanosomes that have been transferred to the cytoplasm of keratinocytes (MN, melanocyte nucleus; MC, melanocyte cytoplasm; K, keratinocytes; G, Golgi apparatus; RER, rough endoplasmic reticulum; *basement membrane at the dermal–epidermal junction; D, dermis) (electron micrograph courtesy of Raymond Boissy).

keratinocytes; (3) the dendrite and keratinocyte plasma membranes fuse, opening a cytoplasmic channel through which the melanosomes pass directly from the melanocyte into the keratinocyte (Jimbow et al., 1999; Marks and Seabra, 2001; Yamamoto and Bhawan, 1994). On incorporation into keratinocytes, melanosomes, which are themselves considered to be specialized lysosomes (Orlow, 1995), interact with lysosomes of the keratinocyte and are sequestered within them forming secondary lysosomes (cf. Chapter 7). As a general rule, melanosomes larger than approximately 1 mm in greatest 65

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diameter are packaged singly in secondary lysosomes, whereas those smaller than 1 mm are sequestered in groups of varying size within a secondary lysosome (Jimbow et al., 1999; Quevedo et al., 1986). Lysosomal hydrolases degrade melanosomes as the keratinocytes move to the epidermal surface. The smaller melanosomes packaged in groups appear to be the most susceptible to lysosomal degradation. The degradation of melanosomes appears to reduce the melanin into ﬁner particles, which may increase its effectiveness in shielding the epidermis from penetration and damage by ultraviolet light (Jimbow et al., 1999). The close relationship that exists between a melanocyte and a small population of keratinocytes with which it maintains contact by its dendrites is formalized in the concept of the epidermal melanin unit (EMU) (Fitzpatrick and Breathnach, 1963). An EMU consists of a melanocyte and the population of keratinocytes that acquires, transports, metabolizes, and disposes of the melanin/melanosomes synthesized within the melanocytes (Quevedo et al., 1974) (Fig. 3.1A and B). It is estimated that an EMU contains approximately 36 viable keratinocytes (Quevedo et al., 1974). Although the uninterrupted horizontal distribution of melanin in the outer corniﬁed layer suggests a functional overlap between EMUs, epidermal pigmentation is largely a mosaic of the individual contributions of millions of EMUs. Just as the unaided human eye fails to resolve the large number of individual dots that constitute a newspaper photograph, the same unaided eye fails to detect that individual EMUs vary in their melanin/melanosome content. Consequently, the brain concludes that skin color is essentially homogeneous. However, under the light microscope, it is evident that melanocytes on the ridges of epidermis that press into the dermis are more melanogenically active than those of interridge areas of epidermis that form a ﬂat interface with the dermis. Not only is more melanin found in the basal melanocytes and keratinocytes of the ridges, the stratum corneum above them often appears to be more heavily pigmented than the interridge corniﬁed layer (Quevedo et al., 1989). This condition is partially evident in Becker’s nevus where there is enlargement of the epidermal ridges and an increase in melanocyte numbers (Quevedo et al., 1989). The freckled epidermis of sun-exposed, fair-skinned, red-haired humans is a classic extreme manifestation of the mosaic nature of EMU structure and function (Fig. 3.2A and B). The clusters of EMUs in freckles produce and harbor abundant melanosomes, resulting in islands of dark epidermis surrounded by a sea of light epidermis composed of EMUs containing weakly melanogenic melanocytes (Quevedo et al., 1986). Melanosomes tend to be largest in African Americans, black Africans, and Australian aborigines and are generally arranged singly within secondary lysosomes of keratinocytes (>1 mm). Melanosomes are smaller (<1 mm) in white individuals of European ancestry, Mongoloid Asians (e.g. Chinese and Japanese ancestry), and certain East Indians; they tend to form groups of two or more in secondary lysosomes (Jimbow et al., 1999; Szabo et al., 1972). 66

A

B Fig. 3.2. Black light photographs of freckled subjects with “carrot red” hair. (A) In this portrait, the population of freckles is so dense that freckles lose their individuality as they increase in size and number and ﬁnally merge with one another in facial areas most prone to sun exposure. (B) Close-up view of discrete freckles on a sun-exposed forearm (photographs courtesy of Dr Robert D. Fleischmann).

When sections of skin are examined under the light microscope, the distribution of melanin within the epidermis correlates with the darkness of the skin. In fair-skinned, white individuals, melanin is scant in the epidermis including the basal layer where only occasional keratinocytes have melanosomal caps shielding their nuclei (Gates and Zimmerman, 1953). When skin sections are treated with ammoniated silver nitrate to darken melanin, greater amounts of melanin are found in all epidermal layers, suggesting that melanosomes may lighten in color because they are under attack by lysosomal hydrolases or passing from an oxidized state to a more reduced one during their ascent within keratinocytes from the basal layer (Pathak and Stratton, 1969; Quevedo et al., 1989) (W. C. Quevedo et al., unpublished observations). In the skin of the dark African Americans, melanosomes are largely concentrated in the basal layer where they form supranuclear apical caps in the germinal keratinocytes (Fig. 3.1A). There are fainter supranuclear melanin caps (Fig. 3.1A) throughout the spinous layer, and scattered melanosomes are found throughout the fully keratinized cells of the stratum corneum. As in white skin, silver nitrate

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

treatment reveals greater amounts of melanin throughout the epidermis of black skin, but its greatest concentration continues to be found in the lower levels of the epidermis (W. C. Quevedo et al., unpublished observations). Along a series of progressively lighter African Americans, the epidermis shows a parallel decrease in melanosomes in all layers as well as an increasingly marked unevenness in the melanin burden of individual keratinocytes (Gates and Zimmerman, 1953). At ﬁrst, the uneven reduction in melanosomes is more pronounced in the corneum and spinous layers than in the basal layer. In more lightly colored African Americans, there are islands of heavy melanin deposits, scattered among lightly pigmented ones, forming a mosaic pattern somewhat similar to freckles in white skin (Gates and Zimmerman, 1953). Corneocytes stripped from the epidermis of human subjects by Scotch tape and treated with silver nitrate to darken melanosomes give a more accurate impression of the numbers of melanosomes reaching the stratum corneum (Goldschmidt and Raymond, 1972). There are large numbers of melanosomes forming a large central cluster in the corneocytes of a very dark individual of African descent. There are few melanosomes and no central clusters in the corneocytes of fairskinned, white individuals. Among individuals of intermediate colors, there is a gradient of melanosome numbers and frequency of clustering paralleling gradually increasing CSC/FSC among the subjects examined (Goldschmidt and Raymond, 1972). The density of melanogenically active (dopa-positive) epidermal melanocyte populations varies between different parts of the human body. The most dense populations are found on the face and genitals and the least dense on the abdomen. Melanocyte populations are distributed symmetrically in the human integument, so that those on the left side match those on the right side. Regardless of skin color, all humans have approximately the same melanocyte densities at corresponding anatomical sites (Szabo, 1967). Therefore, differences in human skin color result from differences in the numbers of melanosomes within EMUs and not from differences in the numbers of EMUs. In addition, the distribution of melanosomes within keratinocytes (single vs. multiple melanosomes in secondary lysosomes) may have some inﬂuence on overall skin color (Minwalla et al., 2001; Szabo et al., 1972; Thong et al., 2003). It is generally considered that regional populations of melanogenically active melanocytes are but one expression of the mosaic nature of the skin established and maintained by a normal developmental program that operates throughout life (Jimbow et al., 1999; Quevedo et al., 1989). Nonetheless, this interpretation has been questioned. It has been pointed out that, with the exception of dopa-positive melanocyte populations in male genitalia (~2000 mm-2), the regional variations in melanocyte numbers parallel the amount of chronic sun exposure that the various parts of the body are likely to have received (Jimbow et al., 1999). It is possible that the high density of dopa-positive melanocyte populations in the male genitalia may be brought about by sex hormones.

In young adults, the habitually exposed face has ~2000 melanocytes mm-2, whereas the generally shielded thigh has ~1000 melanocytes mm-2. As the number of dopa-positive melanocytes increases within skin treated with UVR (Quevedo et al., 1969), it is proposed that repeated exposure to UVR brings about an irreversible elevation of melanogenically active epidermal melanocyte populations (Gilchrest et al., 1979). This would in part explain differences in epidermal melanocyte populations found between individuals. Another reason is that the populations of dopa-positive epidermal melanocytes in human skin decline steadily with advancing age (Quevedo et al., 1969). Therefore, age must be taken into account when comparing melanocyte populations between individuals. Although, at all ages, dopa-positive melanocytes are increased in UVR-treated skin, the populations of melanocytes in irradiated old skin are not as great as in young skin (cf. Chapter 23) (Quevedo et al., 1969). It important to be aware that the sizes of melanocyte populations determined by light microscope cell counts of dopa-incubated skin specimens are substantially smaller than those estimated by electron microscopy (Mishima and Widlan, 1967). As essentially all the studies on melanocyte populations in human skin have involved light microscopy, the regional differences, if any, in the absolute size of melanocyte populations, deﬁned as the total number of dopa-positive and dopanegative melanocytes, remains to be determined. Essentially all estimates of epidermal melanocyte populations have been based on counts of melanocytes in a particular physiological state rather than their absolute numbers. Consequently, the two contrasting interpretations of melanocyte distributions in human skin, developmental programs vs. UVR history, must be given equal status until more is known about the true densities of melanocyte populations.

Pigmentation of the Hair: Follicular Melanocytes In mammals, hair is produced in a hair follicle during a hair growth cycle lasting approximately 21 days in the mouse and as long as several years in humans (see Part II for more details). The hair generated by a hair follicle when properly interpreted is a permanent record of its developmental history. This is particularly valid for its pigmentation. For example, Figure 3.3A is a wholemount of a hair from a uniformly intense black mouse. The eumelanosomes are grouped to form steps of a ladder-like structure when the medullary cells that contain them undergo keratinization. The translucent areas are considered to be “air” spaces that develop in medullary cells when the hair emerges above the skin and dehydrates on exposure to the atmosphere (Roth, 1967). The thin vertical struts supporting the steps of the ladder are melanized cortex cells. The outer cuticle of the hair is rarely pigmented and therefore not evident in this photograph. Figure 3.3B illustrates a dilute black (d/d) phenotype. In this case, the clumping of pigmented melanosomes reduces the amount of pigment available to coat the steps, producing a modest reduction in the intensity of black coat color. Figure 3.3C captures a large mass of melanin pigment in a dilute black (d/d) mouse. The eumelanin mass is 67

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A

A

B

B

C

C

situated near the base of the medullary compartment. Smaller clumps are frequently found in this region near the hair club. Figure 3.4A shows a wholemount preparation of a hair from a Light (Blt/Blt) mouse where the upper region of the hairs displays eumelanin masses that are either whole or large fragments of melanocytes that have been dislodged from the upper hair bulb. As a result, there is an absence of melanin in a long segment of the lower hair shaft. In Figure 3.4B, the hair of a Light mouse contains a single dislodged large melanocyte. Owing to the lack of obscuring melanin in the background, the keratinized framework of the medulla and cortex is clearly evident. In Figure 3.4C, although the hair of mice heterozygous for the Blt gene (not shown) contains pigment clumps, the presence of the pink-eyed gene in the heterozygous state corrects the condition, as revealed by the ladder-like pigment pattern. Figure 3.4D illustrates the failure of the pink-eyed gene to correct pigment clumping caused by the leaden (ln) gene. Each picture tells a different tale about gene action 68

Fig. 3.3. Wholemount preparations of pigmented hairs. Magniﬁcation ¥370. (A) Intense black; (B) Dilute black (d/d); (C) Dilute black (d/d). For descriptions, see text.

D

Fig. 3.4. Wholemount preparations of pigmented hairs. Magniﬁcation ¥370. (A) Light (Blt/Blt); (B) Light (Blt/Blt); (C) Dark sepia (P/p; Blt/b); (D) Pink-eyed leaden (p/p; ln/ln). For descriptions, see text.

(McGrath and Quevedo, 1965). Each hair records the interactions of melanocytes and keratinocytes of the follicular hair bulb as the latter pass the melanocytes on their way toward forming the hair. (For coverage of hair growth and pigmentation in depth, see Part II.) In human hair, the cortex dominates and the medulla is small, so that the most evident pigment in the hair is the vertical streams of melanosomes, reﬂecting their entrapment within elongated spindle-shaped cortical cells. This pattern is particularly evident in hair containing eumelanosomes. In some bright red (pheomelanic)-haired humans, the pigment appears for the most part to be a diffuse yellow rather than granular as found in black or brown hair. In human follicular melanocytes producing melanin chemically deﬁned as pure pheomelanin, the melanosomes are spherical membrane-limited vacuoles containing matrices of microvesicles and amorphous proteinaceous ﬁlaments or lamellae (cf. Chapter 7) (Jimbow et al., 1984; Marks and

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

Seabra, 2001). Pheomelanin is unevenly deposited on the inner matrix at ﬁrst but accumulates to produce a mature melanosome that is highly electron dense. Pheomelanic melanosomes are degraded following transfer to keratinocytes. In ultrastructure, the pheomelanosomes of human hair containing chemically deﬁned pheomelanin are identical to the pheomelanosomes of mice and guinea pigs (Jimbow et al., 1984, 1999; Marks and Seabra, 2001). Mature follicular melanosomes in human subjects with black (eumelanic) hair are black and ellipsoidal. Electrondense eumelanin is deposited on a ﬁlamentous/lamellar matrix of the membrane-limited eumelanosome. As a surface for melanin deposition, small vesiculoglobular bodies also make a contribution to the eumelanosomal matrix (Jimbow et al., 1984; Marks and Seabra, 2001). The majority of red-haired individuals studied by Jimbow et al. (1984) displayed chemically deﬁned “mixed melanogenesis,” a mix of pheomelanin and eumelanin synthesis. Two types of melanocytes were found in their hair bulbs. One produced eumelanosomes, the other pheomelanosomes. The pheomelanocytes also produced a mosaic melanosome incorporating features of both eumelanosomes and pheomelanosomes. The population of eumelanocytes was much smaller than that of pheomelanocytes. On occasion, a keratinocyte was observed to contain both eumelanosomes and pheomelanosomes, indicating that it had been serviced by a eumelanocyte and a pheomelanocyte. Eumelanosomes were singly distributed in keratinocytes, whereas pheomelanosomes were arranged singly or in groups and were highly susceptible to degradation (Jimbow et al., 1984). The vast majority of humans today have black hair. Variation in hair color is largely restricted to white Europeans and their descendants. It is among Europeans that red, blond, brown, and black hair colors are common. Red hair may be found in other ethnic groups but is most frequently associated with a pigmentary disorder such as oculocutaneous albinism (Quevedo et al., 1985). In mice, numerous genes have been shown to inﬂuence eye, skin, or coat color. Their action and interaction have produced a bewildering array of color patterns. The complexity of genetic regulation of melanin pigmentation in mice and also in humans underscores the importance of pigments in the adaptation and evolution of mammals (Hearing and King, 1993; Jackson, 1994; Jackson et al., 1994; Jimbow et al., 1999; Ozeki et al., 1995; Prota et al., 1995; Quevedo et al., 1985; Silvers, 1979; Witkop et al., 1989). In contrast to humans, eumelanosomes and pheomelanosomes have been found within the same follicular melanocyte of agouti mice. They were observed during a shift from eumelanogenesis to pheomelanogenesis (Jimbow et al., 1979; Sakurai et al., 1975). Mice have a juvenile coat of hair that is replaced by an adult one that may differ from it in pigmentation. For example, on some genetic backgrounds, the hair coats of juvenile lethal yellow (Ay/–) mice may appear quite dark (“sooty”) but become clear yellow in adults (Silvers, 1979). Apparently, modiﬁer genes stimulate eumelanin synthesis in hairs pro-

grammed to synthesize pheomelanin speciﬁed by the a-locus, but their ability to do so is restricted to a brief period early in life. Recessive yellow (e/e) mice also exhibit a sooty pheomelanic juvenile pelage that becomes clear yellow in adults (Silvers, 1979). Although the hair coat of light (Blt/Blt) mice is eumelanic throughout life, light mice parallel yellow mice, in that their juvenile pelage is darker, more melanized, than the adult one (Quevedo and Chase, 1958). It is well known that humans with blond or red hair when young may become brunettes with advancing age.

Pigmentation of the Dermis: Dermal Melanocytes Dermal pigmentation results from the presence of melanized melanocytes within this skin layer. Melanocytes synthesize melanosomes that they retain and, possibly, degrade and replace periodically. Although dermal pigmentation is not uncommon in infants of certain human ethnic groups, it is relatively rare in the adult. The classic expression of normal dermal hyperpigmentation is the Mongolian spot or congenital dermal melanocytosis observed in newborns (cf. Chapter 50) (Mosher et al., 1987). The lesions impart a blue–black tint to the skin and range from the typical size of a few centimeters in diameter to very large ones, in some cases extending over the lower back and buttocks. The macroscopic Mongolian spots typically disappear gradually, frequently within a few years. However, persistent Mongolian spots have been found in 3–4% of adult Japanese. The Mongolian spots consist of dermal nests of melanocytes that have been shown to become melanotic at 3 months’ gestation and to lose their melanogenic capacity (dopa reaction) in children from 2 to 10 years of age. Despite the disappearance of the macroscopic spot as a consequence of melanocyte degeneration, inactive dermal melanin-bearing melanocytes have been found throughout life (Kikuchi and Inoue, 1981). The Mongolian spot is evident at birth in 90% of Asian (Mongoloid) and Native American (Amerindian) infants. It is less common in African Americans, black Africans, and European whites (less than 10% in the latter) (Mosher et al., 1987). There are a number of nevi (noncongenital) such as the Nevus of Ota (cf. Chapter 50) that bear some resemblance to the congenital Mongolian spots (Gonzalez-Campora et al., 1994; Mosher et al., 1987). The blue color imparted to the skin by dermal pigmentation requires explanation (R. R. Anderson, in Mosher et al., 1987). When the epidermis is separated from the dermis of the Nevus of Ota and placed on a neutral-colored background, it appears gray or brown in reﬂected light. It appears brown or yellow when observed by transmitted light. In contrast, the pigmented dermis, in the absence of the epidermis, appears blue in reﬂected light. The melanin dispersed in the dermis absorbs incident visible light such that the diffuse reﬂectance in the longer (red) wavelengths is reduced, giving the pigmented sites a blue appearance. However, epidermal melanin does the opposite, eliciting a reduction in diffuse reﬂection at the shorter wavelengths, giving the epidermis a yellow to dark brown tint. As reﬂected radiation from the dermis must reach 69

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the observer by passing through the epidermis, the degree to which dermal melanin appears blue depends on the amount of melanin in the epidermis. Therefore, Mongolian spots with little overlying melanin should appear bluer than those with more epidermal melanin. Possibly, this is why they are so striking in infants before full CSC has been acquired. In addition, based on the contribution that hemoglobin makes to the brightness of blue reﬂectance from dermal melanin in mandrills (see below), the vascular system may have some inﬂuence on the blueness of Mongolian spots in humans. Interference with dermal bluing of the skin by epidermal melanin is suggested by Mosher et al.’s (1987) statement that: “The color [of the dermal melanocytotic nevus of Ota] may be of various hues: black, purple, blue-black, slate-blue, purplish brown and brown . . . Curiously and unexplained, the intensity of the pigmentation may vary daily, and in females during menstruation.” Based on the mandrill, vascular changes bringing more or less blood (hemoglobin) to the spots may produce these rapid color changes (Quevedo et al., 1981).

The Dermal Melanocytes of Other Mammals In wild-type (haired) mice, melanotic dermal melanocytes are commonly found throughout the skin during embryonic development. Melanized dermal melanocytes routinely persist in the relatively glabrous skin (e.g. ear pinnae, tail, and perineum) of the wild-type adult. However, along with epidermal melanocytes, they essentially disappear from the trunk (haircovered) skin during the ﬁrst month of postnatal life (Quevedo et al., 1966). Occasional free dermal melanocytes and dermal perifollicular networks of melanocytes are found in the trunk skin of adult wild-type mice (Quevedo, 1981a). Their frequency of expression is greater in certain mutant varieties of mice. For example, greater numbers of dermal melanocytes persist sprinkled throughout the dermis of adult hairless (hr/hr) mice, suggesting a regulatory defect in a system that normally orchestrates their demise (Quevedo et al., 1995). Compared with wild-type mice, dermal perifollicular networks of melanocytes are also more evident in mice expressing the gene for “dominant alopecia” (Mann, 1969). Well-developed dermal perifollicular melanocytes are found in the hairy trunk skin of the adult Syrian golden (agouti) hamster (reviewed by Quevedo, 1981a). The perifollicular melanocytic networks are associated with the connective tissue sheaths of hair follicles and are located near the sebaceous glands. Topical applications of croton oil, a tumorpromoting agent in which the active ingredient is TPA (12-Otetradecanophorbol-13-acetate), greatly increase the number of perifollicular melanocytic networks, suggesting that many hair follicles have afﬁliated amelanotic dermal melanocytes that are activated and possibly proliferate in response to croton oil (Quevedo, 1981b). Pigmented dermal perifollicular melanocytic networks in normal (untreated) mice are most often associated with specialized tylotrich hair follicles. Tylotrich hair follicles are richly innervated, produce coarse hairs, and function as specialized mechanoreceptors (Straile, 1964). (See Part II for a description of Straile’s model of a 70

cutaneous mosaic of hair follicle groups each surrounding a tylotrich hair follicle and possibly regulated by the nervous system.) Although melanotic perifollicular melanocytes are thought to be absent from the tylotrich follicles of primates including man (Mann, 1969), they are present around the sensory whiskers of at least one primate species, the New World squirrel monkey, Saimiri sciurea (Machida and Perkins, 1967). It is worth emphasizing that the existence of amelanotic perifollicular melanocytes accompanying tylotrich follicles has not been ruled out for primates. As melanocytes are known to synthesize cytokines (Nordlund, 1992), perifollicular melanocytes may be more important for this function than for melanogenesis. If this is the case, either amelanotic or melanotic melanocytes might release cytokines. The tylotrich follicle has been found in the skin of a great variety of mammals, suggesting that it is old in its origin and plays an important role in neurophysiology at the levels of the skin and organism. The mammals possessing tylotrich follicles include duckbilled platypus, spiny anteater, American opossum, European hedgehog, starnose mole, little brown bat, domestic cat, ocelot, tiger, lion, puma, domestic dog, pig, domestic sheep, Japanese deer, house mouse, rat, hamster, guinea pig, porcupine, rabbit, baboon, and a broad assortment of monkeys.

The Interplay of Dermal Melanin and Hemoglobin in Skin Color: The Mandrill (Mandrillus sphinx) One of the most dramatic displays of dermal melanin as a secondary sexual characteristic is in the mandrill (Fox, 1976; Hill, 1955, 1970; Napier and Napier, 1967; Zuckerman and Parkes, 1939). In the adult male of this species, a brilliant red band runs down from the center of the enlarged muzzle from the top of the nose to the base of the nostrils and joins the red lips. On either side of the red nose are brilliant electric blue paranasal swellings, which are cleaved by skin folds. There are tufts of white hair on the cheeks and a yellow to orange beard. The anogenital region of the male roughly matches the “red, white, and blue” pattern of the face. The circumanal region is a brilliant scarlet red and the callosities (thickened skin pads) a pale pink. Immediately lateral to each callosity, the skin is sky blue. Still more laterally, the skin is purplish lilac. The skin extending from the callosities to the penis is blue. The scrotum grades from lilac to pink. There are numerous bright orange scrotal hairs. The penis is pink and the foreskin is scarlet. Although there have been few histological studies of Mandrillus sphinx, the areas characterized by blue coloration are stated to contain abundant dermal melanotic melanocytes with an overlying epidermis devoid of melanin (Fox, 1976). It is this combination of a translucent epidermis with a pigmented dermis that provides the conditions of light absorbance and reﬂectance that produce the blue coloration observed in Mongolian spots. With sexual maturity, major changes in the vascularity occur, resulting in the brilliant blue and scarlet colors of the face and anogenital region. Superﬁcial blood sinusoids develop in the nasal tract of the male

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

mandrill after sexual maturity and become engorged with blood producing the bright red color (Zuckerman and Parkes, 1939). At this time, there is increased blood volume in regions with dermal melanin that produce, depending on the speciﬁcs of vascularity and pigmentation, colors ranging from blue to lilac. In the adult female Mandrillus sphinx, the bright red color characteristic of the male is absent, in part because of the presence of greater amounts of epidermal melanin, which act to obscure dermal blood vessels (Hill, 1955). Nonetheless, in females, the paranasal swellings are a “livid blue” in color and traces of red may be found on the nose. With regard to the anogenital region of the female, there are dark and light areas, but color differentials are in no way comparable to those expressed in the male (Hill, 1955). In male guenons (Ceropithecus spp.), dermal melanocytes in the scrotum contribute a blue color that is more vivid in some species of the genus than in others (Fox, 1976; Napier and Napier, 1967; Wickler, 1967). In free-ranging male mandrills (Mandrillus sphinx), there are positive correlations between testosterone levels and position in the dominance hierarchy. Positive correlations exist between dominance rank (which male wins in agonistic encounters), “levels of plasma testosterone, testicular volume, body weight, and development of secondary sexual characteristics (red and blue sexual skin on the muzzle and rump areas)” (Wickings and Dixson, 1992). Such coloration may be important in sexual selection, for females in some animal species respond to males with the brightest colors, presumably because they signal good health and general ﬁtness. In brief, hemoglobin in blood vessels within the heavily pigmented dermis is required to produce the electric blue coloration found in the male mandrill and, probably, guenons. The combination of changing blood supply and ﬁxed dermal melanin in specialized areas of their skin makes possible transient ﬂashes of bright blue color, the mammalian equivalent of the physiologic color changes brought about by dispersion and aggregation of chromatophore pigments in the lower vertebrates.

Humans Among the Primates The pongids (gorilla, chimpanzee, and orangutan) of today have an extensive covering of hair. Among the early members of the human family descended from pongid ancestors, this coat of hair was reduced but not discarded (Montagna, 1985). Hair was dramatically reduced on all parts of the body except for the scalp, beard, axillae, and pubis, sites at which it played roles in sex-related ornamentation, protecting the skin from damage, in projecting agonistic social signals and, in the case of the axillae, reduction of friction between interfacing skin surfaces (Quevedo et al., 1985). Middle-aged white males acquire gray hair that is largely restricted to the scalp, beard, and chest (Fig. 3.5) (Quevedo et al., 1985). The display of this pattern in more hairy early men would have been quite striking and may have symbolized strength and experience, reducing overt challenges from young males while attracting females. Guthrie’s (1970) life cycle illustration of changing

Fig. 3.5. Composite drawing of graying patterns observed in six white middle-aged males superimposed on a gorilla for artistic convenience. It is evident that the graying of the beard, scalp, and upper chest is intimidating when the length of the hair is exaggerated. Except for the blaze of white on their chests, the humans matched the gorilla (in this illustration) in having pigmented general body hairs. Among the human subjects studied, there appeared to be a strong tendency for the beard to develop streaks of gray initially that gave way to more uniform graying with advancing age.

patterns of the scalp and facial hair of the male suggests that baldness, if early men lived long enough to bald, would have caused a male’s display to shift from a positive signal of virility to a negative one of implied vulnerability. Thus, the tendency for age-dependent male baldness may have devalued the currency of displays originally designed to project male strength and potency. The ability to project dominance signals via coat color in hominids is documented in the saddle-like pattern of gray found on the backs of mature male gorillas (Napier and Napier, 1967). It suggests that displays involving both hair coat and skin color may have formed part of the behavioral program of ancestors common to humans as well as chimpanzees and gorillas, their closest relatives. Recently, there has been renewed interest in hypertrichosis in humans, a condition that, in the extreme, is characterized by a dense covering of body hair sparing only the palms of the hands and soles of the feet. In some cases, although increased, body hair is less dense, ﬁner, and more limited in its distribution. Most types of hypertrichosis are associated with severe developmental abnormalities. Recently, the gene for a form of hypertrichosis that is essentially free of any associated developmental abnormalities has been assigned to the long arm of the X chromosome (Figuera et al., 1995). Congenital generalized hypertrichosis is inherited as an X-linked dominant trait. It causes generalized hypertrichosis in males but patchy hyper71

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trichosis in females. Females are typically heterozygous for the mutant gene and, owing to random X inactivation early in development, the wild-type and mutant alleles are randomly expressed. As a result, a cutaneous mosaic of scattered patches of normal and mutant skin is produced, thereby reducing the severity of hypertrichosis (Figuera et al., 1995). It is possible that the gene mutation that produces hypertrichosis today may be a reversion to a very ancient (“wild-type”) hypertrichosis allele that might have mutated many millions of years ago to precipitate the loss of the dense hair cover characteristic of other primates (Angier, 1995; Figuera et al., 1995; Hall, 1995). It is speculated that the “hypertrichosis locus” may have some additional major function besides regulating hair density and distribution (Angier, 1995). This would account for the persistence of the current so-called “normal” but historically mutant gene that does not perform its ancient function of inducing a dense coat of hair. Periodically, this impotent gene, as far as hair growth is concerned, may undergo back-mutation, leading to a reactivation of the program for hypertrichosis. This model implies that hair reduction early in the evolution of humans or prehumans may have been a very simple genetic event (i.e. a saltation, a jump, producing a new developmental state without gradual change through intermediate forms). With regard to human pigmentation, knowledge of the graying patterns of individuals manifesting extreme congenital generalized hypertrichosis would be particularly interesting, for it might reveal the impact of aging on the display patterns of members of the human lineage prior to the time that the hair coat was reduced historically. With the reduction in hair cover on the appendages and torso in early man, epidermal melanin took on greater signiﬁcance. Like humans, the gorilla and chimpanzee, their closest relatives, have abundant melanogenic epidermal and follicular melanocytes and few dermal melanocytes. Nonetheless, the importance of epidermal pigmentation in humans is still debated. Suggestions as to why melanin is or has been important as a human adaptation include: (1) it absorbs UVR, thus increasing resistance of the skin to sunburn, premature aging, and cancer; (2) it camouﬂages humans in evasion of predators and in hunting; (3) it facilitates thermoregulation by enhancement of absorption of solar radiation or by loss of metabolic heat; (4) it regulates vitamin D synthesis by inﬂuencing penetration of UVR into skin; (5) it protects light-sensitive substances from photodestruction; (6) it interacts with scalp and beard in producing aggressive display patterns; (7) it forms part of a system designed to neutralize the action of noxious chemicals reaching the skin, either directly from the external environment or indirectly from those generated when exogenous agents (e.g. UVR) damage the skin (e.g. free radical compounds) (Nordlund, 1995; Pathak, 1995; Quevedo et al., 1974, 1985). Most, if not all, these suggestions have been challenged, and negative attributes of epidermal melanin have been cited (Hill, 1995; Quevedo et al., 1995). For example, it has been suggested that heavily pigmented humans are more sensitive to frostbite than more lightly pigmented ones. This suggestion has also been questioned (Robins, 1991). 72

Melanin and Human Adaptations to Solar Radiation The effectiveness of melanin as a sunscreen in humans has not gone unchallenged (Kligman, 1995; Nordlund, 1995). This debate is not a new one for, in 1961, Harold Blum ranked its photoprotective value below that of other components of human skin (thickness of the epidermis). In fact, Blum went much further: The widely accepted idea that melanin pigment in human skin protects against sunlight, and that this has bearing upon adaptation to life in the tropics and the distribution of races, [when] examined in terms of its physical and physiological aspects . . . appears to have little merit. It is concluded that whereas the pigment may have a slight adaptive value as regards some aspects of the organism–environment relationship, it may be non-adaptive as regards others; and the respective values may depend upon various complicating factors of the environment. Moreover, the distribution of races according to skin color [if it is adaptive] does not appear to conform well to what would be expected from the spectral distribution of sunlight. (Blum, 1961) While there is logic to Blum’s argument, there is also a fatal ﬂaw, for his bottom line is essentially one of perfectionism rather than relativism in evaluating melanin as an adaptive trait (Blum, 1961, 1969). Relativism holds that an adaptation is not necessarily perfect; it might have both good and bad effects. As small advantages appear to have determined the survival of many lineages in history, as long as the virtues outweigh the liability of a trait, it has a chance to be a winner in the sweepstakes of natural selection. Blum recognizes relativism when he constructs a balance sheet for the positive and negative values of melanin. However, he ignores relativism when he concludes that it is highly questionable whether melanin is a signiﬁcant human adaptation. Failing to ﬁnd a perfect match between melanin’s properties, the worldwide distribution of human skin color, and the problems facing humans in different ecological settings, he assigns melanin a minor role, if any, in the origin, survival, and territorial expansion of the human species. However, based on the numerous possible functions of melanin in humans (Quevedo et al., 1974, 1986, 1995), more than those mentioned by Blum, it is likely that melanin meets a multiplicity of needs simultaneously. Recently, Gilchrest et al. (1996) have demonstrated that increased melanogenesis is stimulated by DNA dimers excised during repair of UVR-induced DNA damage. This intimate link between photorepair of DNA damage and melanogenesis leading to melanin ampliﬁcation indicates that melanin forms an important part of the homeostatic photoprotection mechanism of skin. As different functions of melanin may require different amounts, the level of melanin maintained in human skin, although less or more than the optimal amount for any one function, nonetheless is the best compromise amount for meeting all melanin’s duties while sacriﬁcing as little as possible in effectiveness in any one of them. Therefore, there is an adaptive compromise between contrasting functions of

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

melanin in skin rather than an impossible perfection in all of them. Jacob (1982) has emphasized that evolutionary change results from the tinkering of existing structures and substances into new forms and functions. Consequently, evolution does not produce optimal designs, for it can only work on the materials at hand. The failure to recognize this reality greatly weakens Blum’s position. Whether melanin is or is not a good sunscreen will be determined by thoroughgoing research. However, whatever the outcome of quantitative research, the results will have to be evaluated in the light of a qualitative question: Where would the human species be if melanin was absent from the skin of the early naked human ape evolving in the tropics? The most reasonable answer seems to be: If the emergence of humans was an evolutionary long shot, the odds would have been even longer for humans lacking cutaneous melanin pigmentation. Apart from not considering adaptive compromise, Blum failed to factor in the human life cycle. Among early hominids evolving in the tropics, dark skin color would have been particularly adaptive during early childhood, a time when susceptibility of the still developing skin to solar damage would have been at a maximum. Childhood is also the time when the risk of death from starvation or infectious disease is greatest. It is possible that the adult levels of constitutive skin color (CSC) are primarily the result of natural selection acting on pigment levels during infancy and childhood (Quevedo et al., 1985). In unclad, depigmented, prehistoric humans living in the tropics, sun damage would have been the straw that broke the camel’s back, particularly in children attempting to cope with parasites, infectious diseases, diarrheal dehydration, and malnutrition, conditions that would have been exacerbated by severe sunburn (Quevedo et al., 1985).

The Evolutionary and Biological Basis for Melanin Photoprotection and Phototoxicity Based on the Melanin Content of Human Skin Based on the principle of adaptive compromise, one might envision a scenario for human evolution as follows (Quevedo et al., 1995): 1 The skin of the earliest members of the human family evolving in the solar-inundated continent of Africa was probably hair covered and lightly or darkly pigmented. When reduction in body hair took place, if the skin was lightly pigmented, there would have been strong natural selection for darker skin color, particularly during childhood. Consequently, today’s black Africans are resistant to sunburn and solar-induced skin cancer. They also have the ability to amplify their CSC substantially by increasing the levels of epidermal melanin following exposure to solar radiation, particularly its UVA and UVB components. 2 Melanin distributed throughout the viable and corniﬁed layers of dark pigmented epidermis, based on biological assays, would have blocked the passage of 90% or more of incident UVB and 80% or more of incident UVA to the dermis. Much of the UVB would have been blocked by the stratum corneum (Kaidbey et al., 1979).

3 At low doses of solar radiation, UVA and UVB reaching the viable layers of the epidermis would have generated free radicals subsequently neutralized by general antioxidants and the free radical scavenging redox actions of melanin. The result would be survival and restored functions of epidermal cells. At higher doses, signiﬁcant damage wrought by free radicals in excess of the reducing capacity of the antioxidant defense system would have brought a second line of defense into action. In melanocytes, cytotoxic melanin, its cytotoxic monomers, as well as their cytotoxic precursors released from UVR-damaged melanosomes would exacerbate UVR-induced nuclear damage (DNA strand breaks) and cytoplasmic membrane damage resulting in melanocyte death. Similar events would transpire in UVR-damaged keratinocytes where melanin and its breakdown products would be released from secondary lysosomes (phagolysosomes) and would destroy the functionally impaired cell (Daniels and Johnson, 1974; Quevedo et al., 1995; Riley, 1974). Quevedo et al. (1995) emphasize that: “Melanin/melanogenesis-induced death of photodamaged melanocytes and keratinocytes in vivo may be important in rapidly restoring the essential ‘housekeeping’ functions of sunburned skin and in lowering the potential incidence of skin cancer.” In particular, the removal of damaged stem cells (melanocytes and keratinocytes) of the germinal layer of the epidermis would prevent the cloning of defective cells, which would otherwise chronically undermine essential functions of the epidermis (Riley, 1974; Young, 1987). Quevedo et al. (1995) point out that “[in darkly pigmented humans] the melanin-associated deletion of genetically and functionally compromised melanocytes and keratinocytes is an important factor in the restoration of UVR-disrupted epidermal homeostasis in the short term and in the reduction of precancerous and cancerous lesions in the long term.” The scenario thus far gives substance to melanin adaptations 1 (sunscreen), 5 (photoprotection of vital metabolites), and 7 (detoxiﬁcation of noxious chemicals) listed on p. 72. Melanin adaptations 2 (camouﬂage), 3 (thermoregulation), 4 (regulation of vitamin D synthesis), 6 (aggressive display patterns), and 7 might have forced a level of melanin pigmentation less than optimal for defense of the skin from UVR damage. If childhood was the critical time when melanin exerted its major inﬂuence in depressing mortality in the face of disease and starvation, then photoprotection (melanin adaptations 1, 5, and 7) and possibly photosynthesis of vitamin D (melanin adaptation 4) may have overshadowed all the other postulated functions of melanin. 4 The depigmentation of skin in many human populations following their dispersal from Africa produced a range of colors dominated by individuals with black hair, dark eyes, and moderate levels of constitutive skin color, coupled with marked ability for facultative tanning on exposure to solar radiation. In effect, for most humans, the photoprotection provided by the dark constitutive skin color of Black Africans was retained. Therefore, the same statements made about adaptive compromise in Black Africans holds for most other humans living today. 73

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5 In what is today Western Europe, populations of highly depigmented (fair-skinned) humans evolved with limited ability for facultative darkening in response to solar radiation. The Celtic peoples with red hair, blue eyes, and fair skin are representative of this group. Their melanocytes lack the ability to produce melanin in quantities adequate to provide an effective sunscreen. In addition, melanin/melanogenesis levels are too low to provide effective free radical scavenging or to elicit the death of functionally compromised melanocytes and keratinocytes. As a result, the irradiated skin of such individuals is at greater risk for developing a variety of benign (freckles and keratoses) and malignant (melanoma, basal cell carcinoma, and squamous cell carcinoma) lesions (Quevedo et al., 1995). Considering the negative features of depigmented skin and suggestive evidence that there may have been secondary repigmentation of some depigmented human populations (e.g. Asian Indians), there appears to be no clear adaptive explanation for why reduction in cutaneous melanin should have occurred in the ﬁrst place. To account for geographic/racial differences in skin color, major emphasis has been placed on (1) the requirement for UVR-induced cutaneous synthesis of vitamin D in humans inhabiting latitudes where sunlight is limited and where dietary sources of the vitamin are scarce and/or (2) although controversial, evidence suggesting that black skin is more susceptible to frostbite than white skin. There is no strong consensus of opinion in favor of either one of them (reviewed by Robins, 1991). Perhaps lightening of skin color in certain human populations dispersing out of Africa reﬂected persistence of genetic changes permitted by a reduction in the formerly strong selection for cutaneous melanin in the sundrenched tropics (Quevedo et al., 1985). The resolution of the problems surrounding the current role of the melanin pigmentary system in the skin of humans will have to be solved in humans, for the comparative anatomy of melanin pigmentation elsewhere among the primates is more bewildering than informative.

Cutaneous Melanin Pigmentation of Nonhuman Primates Machida and Perkins (1967) estimated the degree of melanotic pigmentation within the general body skin of 49 species of nonhuman primates. They found that, paralleling rodents, dermal melanocytes in primates are most frequently found in specialized areas of skin, such as the eyelids, lips, nose, and anogenital region. Overall, although some primate species have dense populations of dermal melanocytes in their general body skin, the numbers of dermal melanocytes are reduced in the general body skin compared with specialized skin. Taken as a whole, the more primitive primates, prosimians, have little dermal melanin in their general body skin and, in many species, epidermal melanin is absent as well. Dermal melanin increases in frequency among the Old and New World monkeys but is sharply reduced in humans and their closest relatives, the gorilla and chimpanzee. This brief survey suggests that epidermal and hair follicle pigmentation may be primitive traits dating from the origin of primates and, there74

fore, different from dermal pigmentation, which may be a derived character arising independently in some lineages but not others during their evolutionary divergence. It is important to emphasize that the patterns of epidermal and dermal pigmentation are highly variable in the Old and New World monkeys. Perkins (1975) considers the variation in pigmentation within any group of monkeys to be greatest among the New World monkeys. One might speculate that there has been a reduction in evolutionary selection for epidermal and dermal pigmentation in monkeys, possibly because selection acts only on their exposed surfaces, i.e. their hair coat, and it is adequate for photoprotection. Another possibility is that they live mainly in shaded areas where the hair coat, regardless of density, is sufﬁcient to protect the skin from solar damage. As a result, one ﬁnds a kaleidoscope of cutaneous pigment patterns that, at least in some cases, appear to have no obvious positive or negative adaptive value. The rhesus monkey illustrates the difﬁculty in ﬁnding adaptive explanations for pigmentation in hair-covered skin, for it is not clear how its presence would improve the ability of the organism to survive and reproduce in its native habitat. In the rhesus monkey, the hair-covered general body skin, but not the hair, is pigmented in a piebald (spotted) fashion. The epidermis lacks melanogenic melanocytes, but they can be induced by treatment with UVR (Erickson, 1976; Montagna et al., 1964; Yun and Montagna, 1966). Patches of dermal melanocytes are scattered throughout the skin. Within the pigmented patches, dermal melanocytes are frequently observed to be wrapped around blood vessels. Although UVR stimulates epidermal pigmentation, it does not inﬂuence the activity or distribution of dermal melanocytes (Erickson, 1976; Montagna et al., 1964; Yun and Montagna, 1966). UVR has also been demonstrated to increase the number of melanogenic melanocytes in the epidermis of the potto, Perodicticus potto (Montagna, 1967). In view of the ability of melanocytes to synthesize cytoactive substances, it is possible that melanocytes enveloping blood vessels might play a role in skin physiology unrelated to melanogenesis. The same can be said for melanocytes associated with sebaceous glands and the perifollicular networks of dermal melanocytes. James J. Nordlund (1992) is to be credited for emphasizing the roles played by melanocytes in processes other than those associated with melanin. Mammalian melanocytes are considered to be important modulators of inﬂammation and have been demonstrated to synthesize interleukin (IL)-1 and transforming growth factor (TGF)-a and other growth factors and cytokines (Nordlund, 1995). It is important to keep in mind the diversity of functions that melanocytes may perform in the hairy skin of primates when attempting to interpret their often bizarre distribution patterns. The cases in which pigmentary explanations for purpose in melanocyte patterns utterly fail may be ones where closer scrutiny reveals heretofore unknown functions of melanocytes, or familiar ones seen in a different context. From this perspective, biologists should not be frustrated by evolutionary paradoxes in pigmentation but rather they should be delighted.

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

The Hair Coat as the Interface between the Mammal and its Environment Coat color patterns are considered to be important in protective (concealing) coloration, sexual attraction, warning displays, and thermoregulation. Although the distribution of epidermal and dermal melanocytes shows signiﬁcant arbitrariness in the skin of many mammals with dense coverings of hair, the color pattern of the hair coat does not. It appears to be produced by developmental genetic mechanisms that have been closely scrutinized by natural selection. However, nongenetic maternal environmental factors operating during gestation may have some inﬂuences on coat color patterns (Caro and Durant, 1991). African cheetahs (Acinonyx jubatus) are dark spotted over most of their bodies with the exception of the apex of the tail where there are stripes. The pattern of tail stripes of sibling cheetahs is very similar and distinguishable from the array of tail stripes on their mother. The pigment patterns of siblings are more alike than they are similar to those of nonsiblings of equal age. It has been hypothesized that “environmental factors in utero are responsible for (pattern) differences in cheetahs in combination with maternal or paternal genetic inﬂuences,” thus producing the greater phenotypic coat color variation observed in cheetah populations than expected in a species with very high levels of genetic homozygosity. It is possible that coat color patterns may directly inﬂuence bonding or are markers of other developmental events that control bonding. Male cheetahs may form coalitions consisting solely or predominantly of brothers. Such coalitions have better odds of holding territories favoring hunting and reproductive success (Caro and Durant, 1991). Individual and geographic variation in pigmentation patterns of the harbor porpoise, Phocoena phocoena, suggests that melanin pigmentation may play a role similar to that in the cheetah in recognition between individual porpoises of a group. However, the variation in color patterns of geographically isolated groups of porpoises is insufﬁcient to establish taxonomic distinctions between them (Koopman and Gaskin, 1994). Unlike the symmetrical pigmentation patterns in the cheetah, those of Phocoena phocoena are asymmetrical (Koopman and Gaskin, 1994). Mammals may escape from constraining developmental rigidity of pigmentation in another way. In adapting to temperate habitats, mammals such as the snowshoe hare have seasonal changes in hair color such that hair grown in the spring is brownish and that grown in the fall is white (Keogh, 1969). In response to changing photoperiods, hormones trigger the appropriate melanocyte responses (Rust, 1965; Rust and Meyer, 1968). However, the changes brought about are economy minded, for the whitening of the coat involves the growth of overhairs that are white (pigmentless) at their exposed tips but dark in their concealed bases. All the concealed undercoat of insulating hairs remains pigmented regardless of season. Such selectivity in the types of hairs participating in seasonal color changes and the mathematical precision with which hairs are partially pigmented suggest a

tightknit relationship between the neuroendocrine system and the skin (Rust, 1965; Rust and Meyer, 1968). The thin winter coating of white suggests that the seasonal color changes in the snowshoe hare are important in concealing it from predators. The permanent white coat of the polar bear is considered to be important in concealing it from prey when hunting on ice and snow. In summer, in some parts of the world (e.g. the Hudson Bay area, Canada), with the melt of sea ice, it is not uncommon for polar bears to be restricted to land temporarily devoid of ice and snow (Nowak, 1991). Therefore, at least for some populations of polar bears, the white coat may prove a seasonal disadvantage for, rather than being concealing, it calls attention to its owner. This disadvantage may be partly reduced by the polar bear’s omnivorous lifestyle. Seals are the major source of food, particularly in winter. Depending on location, when feeding on seals is not an option in summer and fall, polar bears feed on sea birds and their eggs plus berries and other vegetation. In the absence of snow and ice, its coat color may make it more conspicuous to prey and thus prove a disadvantage. On occasion, polar bears may eat the carcasses of stranded marine mammals, small land mammals, reindeer, and ﬁsh. Nonetheless, a life without seals is a difﬁcult one. In the James Bay region of Canada from July through December, the polar bear lives off stored fat to a signiﬁcant extent, hanging on until the sea ice returns. This suggests that the coat color of polar bears has real advantages and disadvantages, with the former greater than the latter when it comes to feeding (Nowak, 1991). A competing hypothesis is that the white coat and darkly pigmented epidermis of the polar bear are associated with thermoregulation (Nowak, 1991). Possibly, the true explanation for the evolution of the polar bear’s coat is that it was selected for both functions. The agouti (A) coat of feral house mice, Mus musculus, appears to be associated with concealment from predators, as do the light and dark coats of the pocket mouse matching light (sandy) and dark (lava) backgrounds in New Mexico (Cott, 1957). The agouti pattern of coat color in house mice arises from a complex array of hairs that differ from each other in shape as well as the color that they display, thus rendering their bearers less visible, safer, when in harmony with the background coloration of their habitat. For further details about the wild-type coloration of mice, see Part II. The lion cub has dark spots and dark rings on its tail; the adult has not. Presumably, the disruptive coloration of the cub favors concealment in the grassy landscapes on which they are reared and/or possibly identiﬁes them as juveniles, protecting them from adult aggression. The traditional explanation of the alternating light and dark stripes (hair color reinforced by skin color) of the zebra is that they produce disruptive coloration, making it particularly difﬁcult to detect zebras on the African savannahs at dusk when risk of attack is greatest (Cott, 1957). Alternatively, when zebras are in herds, their swift movement may cause a blurring of individual outlines, momentarily distracting their predators (MacClintock and Mochi, 1976). However, these 75

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interpretations have been questioned (MacClintock and Mochi, 1976; Owen, 1980). One suggestion is that the color pattern of the zebra disorients biting ﬂies, decreasing their success (Waage, 1981). In addition, it is possible that zebra stripes, because they vary between individuals, may facilitate intraspeciﬁc recognition as well as membership in a particular herd, a role comparable to that already suggested for color pattern in cheetahs (MacClintock and Mochi, 1976). A variety of evidence suggests that zebras are black with white stripes produced by the gradual evolutionary enlargement and coalescence of small white ﬂecks in the descendants of their darkcoated ancestors (MacClintock and Mochi, 1976). Based on his experiments, Waage (1981) concludes that black-bodied zebras would be more prone to ﬂy bites than their striped counterparts.

The Coat Colors of Juvenile Mammals: A Time to Learn and Play Among mammals, hair color patterns often change between infancy, adolescence, and adulthood in association with the different roles played and problems faced at different stages of the life cycle. The young chimpanzee has a patch of white hair on its rump that elicits tolerance in its misbehavior toward the powerful adults that surround it (Napier and Napier, 1985). The young of many monkey species are dramatically different in color from the adult. The dusky leaf monkey (Presbytis obscura) as an adult wears a dark coat of hair whereas the infant wears one that is apricot in color (Napier and Napier, 1985). In some human populations, hair frequently darkens with age. In these cases, lighter hair color along with other juvenile traits may have some inﬂuence as a releaser of empathy directed by human adults toward their young. Clearly, many social primates emphasize in their coat colors that youth is a time for learning and support by the community. In primates other than man, particularly in the Old and New World monkeys, there is generally a dense covering of body hair, sparing only certain regions such as the face and anogenital region. It is in many of these highly social animals that there is an intricate pattern of glabrous skin pigmentation framed by hairs of various hues. It is clear that these faces are intended to be seen and to transmit information. The associations of tufts of hairs of different colors together with patterned facial skin pigmentation indicates that this complex of features is a deﬁning element of certain species and has been shaped in ﬁne detail by evolutionary mechanisms (Napier and Napier, 1985). In some primates, the color of the hair coat is less important than what is on it. In South American sloths, the coat is covered by algae that impart a green color that camouﬂages the sloths as they hang from tree limbs (Cott, 1957; Owen, 1980). According to Nowak (1991), “the color [of the polar bear] is pure white following the molt but may become yellowish in the summer, possibly because of oxidation by the sun. The pelage sometimes appears grey or almost brown, depending on season or light conditions.” The mechanisms 76

involved are not stated. It is possible that the variations in hair coloration result from the oxidation of lipids of sebaceous gland origin that coat the hair. In some cases, the off-white color of the polar bear might render it less visible, particularly on snowless landscapes and on snow under certain lighting and weather conditions. Although a start has been made (Vrieling et al., 1994), the developmental mechanisms involved in establishing the complex coat color patterns of mammals have not been identiﬁed. A number of different mathematical models have been proposed but have not as yet been attached to real events transpiring within the developing skin (Nagorcka and Mooney, 1992; Pearson, 1993; Quevedo et al., 1986).

Radicalism and Conservatism in the Evolution of the Mammalian Cutaneous Pigmentary System from that of Reptiles The evolution of mammals from reptiles did not occur in one big step. It took place gradually; the traits that deﬁne mammals (warm-bloodedness, diaphragm, hair, etc.) accumulated and became more efﬁcient over considerable periods of evolutionary time. These events transpired within a lineage known as the mammal-like (synapsid) reptiles (Kemp, 1982). The innovative adaptations of mammal-like reptiles to changing environments were put to good use by the mammals, their gift to posterity. Following the demise of the dinosaurs, the adaptive traits of mammals facilitated an evolutionary increase not only in the number but also in the diversity of species. More precisely, the ancestral diminutive mammals of limited diversity rapidly produced many divergent lineages that, in turn, generated a great variety of specialized descendants capable not only of ﬁlling the habitats vacated by the ruling reptiles but also equipped to enter new habitats as well. The skin and its appendages played an important role in determining the ecological success of mammals. The evolutionary changes in the skin associated with the reptile– mammalian transition were complicated ones that tested the relevance of each skin component to emergent new functions. Some components did not stand the test and disappeared; others became less prominent. In the case of the dermal chromatophores, the instruments of change in vertebrates other than the birds and mammals, the cutaneous xanthophores and iridophores presumably disappeared from advanced mammallike reptiles while their dermal melanophores were replaced by smaller numbers of melanocytes lacking the ability for rapid, reversible movement of melanosomes between their cell bodies and dendrites (Bagnara and Hadley, 1973; Oliphant et al., 1992). Only epidermal melanocytes prospered by the evolutionary test. They became the major source of skin pigment in mammals. They retained their ability to transfer melanosomes to keratinocytes but, like the dermal melanocytes, they lost the ability for rapidly aggregating or dispersing melanosomes intracellularly in response to endocrine and neural stimulation (Klaus and Snell, 1967; Reams et al., 1968). In reptiles, all the chromatophores are thought to be derived

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

from pluripotential stem cells of neural crest origin, the speciﬁc differentiated fate of which (melanophore, xanthophore, iridophore) is determined by the particular cutaneous site that each stem cell becomes lodged in during embryonic development (Bagnara et al., 1979). Consequently, major changes in regulation of cutaneous development were indispensable for the evolutionary emergence of mammalian pigmentary patterns produced by epidermal and follicular melanocytes (Quevedo and Holstein, 1992). Three major events in the reorganization of the skin among mammal-like reptiles were: (1) the loss of epidermal scales from the body surface; (2) the development of hair; and (3) the modiﬁcation of the molecular biological pathway for epidermal keratinization, which produced new patterns of molting in those mammal-like reptiles approaching full status as mammals. In the ancestors of mammals, as in their descendants, molting became increasingly identiﬁed with periodic shedding of hair rather than the shedding of corniﬁed epidermis. The loss of epidermal scales resulted from modiﬁcations in epidermal keratinization. Their disappearance over most of the body erased or respeciﬁed a complex mosaic of morphogenetic ﬁelds formerly embodied in the epidermal scales, the intervening scale hinges of which had served as the “grout” that held the mosaic together. This change was not a minor one, for the scales had imposed order on the skin, as evidenced by the intricate associations of diverse dermal chromatophores within each scale territory (Whimster, 1965, 1971). In mammalian skin, there is evidence of a mosaic consisting of repeated “vertical units” containing epidermal, neural, and vascular elements arranged around a sensory tylotrich hair follicle (Maderson, 1972; Straile, 1969). These morphogenetic units may reﬂect in some highly derived way the morphogenetic ﬁelds of scale-covered skin in ancestral reptiles. In reptilian lineages other than that of the mammal-like reptiles, species-speciﬁc skin color patterns continued to be produced by the programming of groups of adjacent scales to make coordinated site-speciﬁc skin displays of coloration. The “scale color display units,” like the colored tiles in ceramic mosaics fashioned by human artisans, collectively gave rise to a grosser distinctive pattern of value in a sighted world. In the case of reptiles, the pattern concealed its bearer, attracted conspeciﬁcs, and repelled potential predators including conspeciﬁcs (Whimster, 1965, 1971). These behavior-triggering functions of pigment patterns emerged when the observing animal contemplated the whole integument of the object of its attention rather than its individual component parts (scales). The loss of scales in advanced mammal-like reptiles was presumably precipitated by events associated with the origin of hair and almost certainly destabilized the reptilian-style dermal pigmentary system from a developmental point of view. In addition, an increasingly dense covering of hair exacerbated the stress imposed by scale loss on the dermal pigmentary system by making dermal chromatophores largely irrelevant in photoprotection and adaptive color changes. Maderson (1972) has outlined a plausible scenario for the

origin of hair and loss of scales in mammal-like reptiles. It centers on the premise that hair ﬁrst arose as a mechanoreceptor that provided mammal-like reptiles with information about the position of their scales and about external objects in contact with their skin. Accordingly, the insulating function of hair must have been a secondary rather than a primary adaptation. The ﬁrst hairs were limited in number and distributed diffusely over the body. The tylotrich hair follicles that serve as mechanoreceptors and their companion hairs show a special distribution in the dorsal skin, which suggests that they arose in the scale hinges and were separated by scales at some time in the remote past. Maderson (1972) suggests that the proliferation of hair follicles to amplify monitoring of the environment (e.g. sensing the air currents passing across the skin) would result in their invasion and destruction of scales. Accompanying the spread of hair follicles would be an altered pattern of epidermal keratinization featuring keratohyalin granules in the superﬁcial epidermis. Maderson hypothesizes that “hair” began as a simple column of epidermal cells but gradually evolved into the multilayered hair follicle of today, a structure signiﬁcantly governed by interactions of a dermal papilla with germinal keratinocytes of the hair follicles (Cotsarelis et al., 1990). The tylotrich hair follicle, with its encircling networks of nerves, is thought to mirror features of the advanced prototypic hairs of progressive mammal-like reptiles. The broad distribution of the tylotrich hair follicle in mammals from duckbilled platypus to humans suggests that it was an early component of the hair coat and in all probability was present in mammal– reptiles (Maderson, 1972; Mann, 1968, 1969) (see Part II for coverage of the embryology of hair formation). By this stage in the evolution of mammal-like reptiles, a sophisticated behavioral pattern for thermoregulation would have evolved so that shifts made in body position in response to air currents would have facilitated appropriate heat gain or loss. Sustained selection for a thickened hair coat to enhance mechanoreception was eventually co-opted to provide insulation in support of thermoregulation (Maderson, 1972). Of the pigment cells in the skin, epidermal melanocytes would have retained an adaptive melanin-based purpose in animals covered with hair. They were required to pigment the hair as well as any areas of skin epidermis that was devoid of hair cover. In addition, many mammals bask in the sun, accumulating heat by exposing their ventral surfaces where hair density is lowest. Reptiles habitually sunbathe. A pigmented epidermis in an early reptile–mammal may have facilitated heat absorption while preventing sunburn of the skin during basking (Hamilton, 1973). Such behaviors reduced the requirement for feeding (caloric intake) in metabolic support of thermoregulation and may have been very important during the transition from exothermy to endothermy. Therefore, the presence of a pigmented epidermis sometimes accompanied by a pigmented dermis in skin exposed to solar radiation was adaptive and thus a target for natural selection leading to developmental genetic programs that assured appropriate 77

CHAPTER 3

embryonic deployment of cutaneous melanocytes. In polar bears, for example, the combination of white fur and melanized (black) epidermis is thought to trap and transfer substantial amounts of solar heat into the blood vessels of the skin. The skin of mammals was programmed regionally in order to specify the presence or absence of differentiated melanocytes in the epidermis and/or dermis in keeping with the specialized functions of the particular site. For example, in the trunk skin of mice, differentiated melanocytes are present in the epidermis, dermis, and hair follicles at birth. Many of the epidermal melanocytes enter the hair follicles and survive. However, numerous epidermal melanocytes do not. The epidermal melanocytes that do not enter hair follicles and dermal melanocytes disappear almost completely during the ﬁrst month of life (Quevedo et al., 1966). In contrast, melanocytes are retained in the epidermis and dermis of essentially glabrous skin such as the perineum, tail, and ear pinnae. The persistence of melanocytes in this case may aid concealment of feral mice as well as play some role in photoprotection and courtship of mice in the wild.

Part II: The Development of the Melanin Pigmentary System and the Molecular Genetic Systems that Regulate It Introduction Thus far, the major focus of this chapter has been threefold: the structure and function of the melanin pigmentary system (MPS) of mammals; the great variety of melanin pigment patterns found in the integument of mammals; and the adaptive signiﬁcance of the MPS during millions of years of mammalian evolution. On the short scale of time, knowledge of the mechanism by which such patterns are laid out during embryonic development is equally as important as the long-term events that shape them. A substantial number of the specialized chapters in this volume are dedicated to exploring in depth the biochemistry, developmental biology, molecular biology, and cellular biology of the MPS that emerges during embryogenesis and functions throughout the life of the organism. By way of an introduction to these chapters, what follows is a brief integrated account of the genetic program that regulates the development of the MPS in the mouse. It is our hope that it will be of assistance to readers without a background in molecular and developmental biology. For readers desiring additional coverage in greater depth, the links between this introduction to development and the matching more technical chapters are listed at the end of this account. To begin with, the development of the MPS is described to establish a context in which to discuss the mechanisms that regulate its origin and function. Based on the large number of hair color mutants known and preserved in laboratory mice, their developmental genetics of 78

pigmentation is particularly well understood. Historically, the development of the wild-type color pattern of the laboratory mouse has been treated as the “standard pattern” from which all other color mutations in laboratory mice have departed. The account of MPS development that follows adheres to this tradition. It is noteworthy that numerous pigment gene homologies have been found between mice and humans.

The Development of the Hair Coat of Wild-type Mice A brief account of the development of the hair coat in mice is in order to establish a structural context within which to discuss regulatory mechanisms related to pigmentation of the hair. Embryonic development of hair follicles is tightly controlled by numerous genes that encode growth factors. In addition to regulating epidermal and dermal cells directly related to hair growth, growth factors, by chance or necessity, may also beneﬁt the invading melanocytes that are of little use to the hair follicle, but of great importance in the survival of the organism as a whole. The hair coat is complex in its structure. According to Chase and Rauch (1950), the dorsal pelage of the mouse consists of at least four distinct types of hairs divisible into two groups: the long overhairs and the shorter underhairs. The overhairs consist of the monotrichs (1.6% of hair coat), awls (8.7%), and auchenes (7.8%). The underhairs, called zigzags, make up 81.9% of the dorsal pelage. In wild-type (agouti) mice, the overhairs differ in the frequency with which they express the agouti band, a band of yellow pigment that divides the otherwise uniformly eumelanic black hair (except for a short portion of the tip and base that lacks pigment) into a hair shaft that contains a black eumelanic band near the tip followed by a yellow pheomelanic band and a long basal eumelanic segment accounting for much of the remaining hair shaft. Zigzag hairs invariably have an agouti band; auchenes usually, but not invariably, have an agouti band; awls lack agouti bands more frequently than auchenes; and the monotrichs invariably lack them (Galbraith, 1964, 1969). In mice, the mix of black hairs and yellow-banded hairs produces a hair coat in which blackness and yellowness are muted to an indeﬁnite intermediate overall color. Hundreds to thousands of tiny ﬂecks of yellow and black may serve as camouﬂage and confuse the eyes of potential predators as well as the human eye so that one mouse can appear dark grayish, brownish, or blackish and inconspicuous depending on the background and ambient light.

Embryonic Establishment of the Hair Coat in Mice The initiation of the growth of hair follicles begins on the 14th day post coitus (dpc) in the mouse and continues until slightly before birth (Mann, cited in Claxton, 1966) or slightly after

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

birth (Claxton, 1966). Its earliest stages are dependent on exchanges of signals between dermal mesenchyme cells (precursor dermal papilla) and overlying foci of epidermal cells (keratinocytes). In response, epidermal cells crowd together, with the precursor dermal papilla condensing beneath them. They are separated by a basement membrane (basal lamina). The crowded keratinocytes sink slightly into the dermis producing a node (bud), the collective cells of which are designated as the initial hair follicle germ. In later hair generations, as discussed below, the equivalents of the initial hair germ and initial dermal papilla interact to induce the growth of a new hair (Hardy, 1992; Lavker et al., 1999). In the mouse embryo, signals from the initial dermal papilla are considered to induce the cells of the epidermal node (bud) to grow into the dermis and elongate into a peg, a cellular cylindrical rod-like structure. In doing so, the solid cylindrical rod (peg) of epidermal cells (keratinocytes) with its former hair germ leading grows over the dermal papilla producing a hair (follicle) bulb that takes the lead as the peg extends deeply into the dermis (Figs 3.6A and 3.7) (Chase et al., 1951). The follicle is now characterized by the hair bulb and two other bulges,

the presumptive sebaceous gland and, slightly below it, a bulge of the outer root sheath (ORS) to which the arrector pili muscle attaches (Fig. 3.7). The ORS bulge is thought to be a reservoir of stem keratinocytes and possibly melanocytes (Akiyama et al., 1995). The permanent (upper) hair follicle ends slightly below the ORS bulge. The part of the hair follicle below the ORS bulge is temporary and is destroyed during each hair growth cycle only to be recreated by the next one. Within the hair bulb of the lower follicle, the basement membrane separates the dermal papilla not only from the surrounding keratinocytes but also from the migratory melanocytes that have invaded the hair follicle (Fig. 3.6B). The keratinocytes of the bulb form a matrix. Within the matrix, the keratinocytes located in approximately the lower half of the bulbous portion of the lower hair follicle dominate in mitotic activity and create a pressure for upward movement of keratinocytes toward the skin surface. In pigmented mice, keratinocytes destined to form the hair acquire melanosomes when they encounter melanocytes adjacent to the dermal papilla in the upper hair bulb. During their upward movement, keratinocytes are organized into concentric channels

K

G

MC

MN DP

G

A

K

B

Fig. 3.6. Vertical sections of hair follicles plucked from a human scalp. (A) Incubation of a follicle in “dopa-reagent” to enhance melanin synthesis decisively distinguishes dendritic melanocytes (arrows) from keratinocytes in the upper hair bulb. Note that the melanocytes reside at the junction of the dermal papilla (DP) and follicular keratinocytes. (B) Ultrastructure of a follicular dendritic melanocyte in intimate contact with several keratinocytes. Numerous melanosomes at formative stages ranging from unmelanized (Stages I and II) (arrows) to fully melanized (Stage IV) populate the melanocyte cytoplasm. Melanized melanosomes are also present in the keratinocytes (arrowheads). (MN, melanocyte nucleus; MC, melanocyte cytoplasm; G, Golgi apparatus; open arrow, melanocyte dendrite; K, keratinocyte) (both micrographs courtesy of Raymond Boissy).

79

CHAPTER 3 Telogen

Anagen I

Anagen II

Anagen III

Epidermis Dermis

Adipose layer

Permanent follicle

Sebaceous gland

Outer root Arrector pili Sheath bulge muscle Temporary follicle

Anagen IV Anagen V Anagen VI Catagen

Dermal papilla

within which the ascending cells meet substantially different developmental fates (Chase et al., 1951; Lavker et al., 1999). Within the solid column of keratinocytes forming the cellular follicular peg above the hair bulb, central cells die and decompose, providing a space within which the upward movement of two major channels of cells, each divisible into subchannels, can take place. Within the most central channel, there is a transformation of streams of medulla, cortex, and cuticle cells into a hair shaft. It grows through the channel in the follicular rod (peg) and out of the hair canal, an opening in the epidermis. This pattern of development not only permits the hair to display itself to the sighted world beyond the skin, but also aids the organism in keeping warm and capable of sensing features of its environment everywhere in the integument that encloses its body [see Straile and Mann (1972) on touch receptors below]. During hair growth, the upper matrix cells of the hair bulb are the ﬁrst cells incorporated into the hair shaft as well as the inner root sheath (IRS) that surrounds the hair shaft. The IRS consists of three layers: an outer Henle’s layer, middle Huxley’s layer, and inner cuticle layer. Although the IRS keratinizes and ascends with the hair shaft toward the skin surface, it stops short and sheds itself into the hair canal near the duct of the sebaceous gland. The outer root sheath (ORS), which surrounds the IRS, although of epidermal origin, is maintained by the epidermis with which it is contiguous. Completing the layers of the hair follicle is the connective tissue sheath of dermal origin that surrounds the ORS of the follicle. It con80

Fig. 3.7. Critical stages in the growth cycle of mouse hair follicles, courtesy of Dr William Montagna (slightly modiﬁed). Left: an actively growing hair follicle illustrating some of its important features. Right: illustration of the stages of the hair growth cycle: telogen (quiescent period), anagen [period of active follicle and hair growth (six substages characterize anagen)], and catagen characterized by the cessation of hair growth and regression of the hair follicle to the quiescent state.

sists of three layers: the inner basement membrane/basal lamina and an intermediate layer of collagen. The outer component of the connective tissue sheath is cellular, composed of ﬁbroblasts and macrophages, and in contact with the papillary area adjacent to the dermal papilla (Lavker et al., 1999; Parakkal and Alexander, 1972). Once the growth of hair is completed, mitotic activity in the bulb matrix is terminated. The hair follicle tends to “deconstruct” itself in the sense that melanocytes stop synthesizing melanosomes and die by apoptosis, a complicated process by which a cell destroys itself (Lavker et al., 1999). The growth of the hair shaft stops and its base is left unpigmented. However, cells that have started to differentiate continue to do so and migrate upward to form the last part of the hair shaft. The hair shaft forms a brush-like ending taking on the appearance of a club fashioned by modiﬁed cortical cells that ascend with it. Once the hair club has been completed and surrounded by transformed ORS keratinocytes (germ cells) that anchor the club hair in the permanent follicle, the lower hair follicle is essentially reabsorbed. The death of matrix keratinocytes is thought to be brought about by terminal differentiation. Other processes such as heterophagy have been suggested as factors in the demolition of the lower hair follicle (Lindner et al., 1997). The dermal papilla moves upward stopping just below the hair germ of the now quiescent hair follicle. During the resting stage of the hair growth cycle, the conﬁguration of the hair germ and dermal papilla is reminiscent of their association at the start of hair follicle formation in the embryo. They are still separated by a basement mem-

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

brane (Chase et al., 1951; Hardy, 1992; Lavker et al., 1999; Parakkal and Alexander, 1972). The recurrent hair growth cycle in mammals consists of three major stages: (1) anagen, during which the hair shaft is formed by the hair follicle; (2) catagen, the transitional stage between anagen and telogen, during which the hair follicle regresses to the telogen conﬁguration; and (3) telogen, (“resting stage”), beginning after the growth of the hair has been completed and a basal club is formed that anchors it in the upper hair follicle (Fig. 3.7) (Chase et al., 1951; Montagna and Parakkal, 1974; Parakkal and Alexander, 1972). All hair growth cycles that follow the ﬁrst one also depend on signaling between the dermal papilla and hair germ. There is some debate as to the source of matrix keratinocytes for each generation. Challenging the concept of the traditional self-perpetuating hair germ as the sole source of matrix keratinocytes is evidence that the matrix keratinocytes originate in the bulge of the ORS, which may also be the source of the cells that form the hair germ for each generation (Chase et al., 1951; Cotsarelis et al., 1990; Parakkal and Alexander, 1972). Like the mouse, the human upper hair follicle is thought to contain a population of stem keratinocytes, which implies that the mouse model for hair growth applies to humans as well (Lavker et al., 1999; Moll, 1995; Yang et al., 1993). A variety of evidence suggests that the melanocytes that pigment hair in one growth cycle are not involved in the pigmentation of the next. First, in light (Blt/-) mice and silver (si/si) mice, melanocytes start to die approximately midway in anagen, resulting in their being uprooted and incorporated into the developing hair shaft (see Figs 3.3 and 3.4) (Quevedo and Chase, 1958; Quevedo et al., 1994). As a consequence, all melanocytes are physically removed from the hair bulb and disposed of in the hair shaft. The hair continues to grow in the absence of melanocytes for approximately a week, so that it is pigmented at its tip and pigment free in much of its base. Nonetheless, the hair bulbs of the next generation have what appears to be a full complement of melanocytes that ultimately meet the same fate as their predecessors. As noted above, during catagen in wild-type mice, both melanocytes and keratinocytes of the temporary region of the hair follicle die, suggesting that melanocytes for the next hair generation come from another source. The presence of amelanotic melanocytes within hair germs of resting follicles was inferred when melanogenic (DOPA-positive) dendritic melanocytes were found following treatment of pigmented mice with multiple small doses of X-rays that stimulate melanogenesis (Quevedo and Isherwood, 1958). Electron microscopy subsequently revealed amelanotic melanocytes containing immature melanosomes (Sugiyama et al., 1979). In addition to their presence in the hair germ, cells in the bulge of the ORS have not only been determined to be stem keratinocytes, but others have been identiﬁed as melanocytes by melanocyte-speciﬁc antibodies (Akiyama et al., 1995). Therefore, although stem melanocytes and keratinocytes in the traditional hair germ and/or bulge might provide a new complement of matrix keratinocytes and melanocytes for each hair growth cycle, it

seems most likely that the ORS is the sole source for both, particularly as the germ cells for each hair follicle generation are apparently derived from the ORS (Chase et al., 1951; Cotsarelis et al., 1990; Parakkal and Alexander, 1972). Recently, Sharov et al. (2004) published a brief account of their work suggesting that, in TRP2-LaxZ transgenic mice, melanocytes incorporated in the follicle germs of “resting” (telogen) hair follicles originate from more than one source. Not only are they derived from amelanotic stem cells of the ORS bulge but also from other ORS amelanotic melanocytes that escape the widespread cell death that occurs within hair follicles during catagen. The latter ORS melanocytes are thought to do so by retreating upward in the regressing, surviving portion of the hair follicle (Sharov et al., 2004). The technology employed in this study is more sophisticated than previous ones dealing with the same topic. The details of the second generation of hair growth, with some exceptions, are comparable to those of the ﬁrst generation. Two examples of the exceptions are, ﬁrst, that the hair germ and dermal papilla are in position to initiate the second hair generation as a legacy of the ﬁrst hair generation. Second, the hair of the second generation exits the skin via the slightly altered transepidermal channel established for that of the ﬁrst generation. The ﬁrst hair shaft may be shed or remain anchored in the follicle so that two or more shafts emerge from one follicular oriﬁce (Chase and Silver, 1969). Embryogenesis of the hair follicle becomes even more complex when viewed populationally. Hair follicles are not distributed randomly in the skin for they are arranged in linear bands in the adult mouse. However, in neonatal mice, they form discrete groups consisting of hair follicles that produce overhairs (monotrichs, awls, and auchenes) and underhairs (zigzags) (Claxton, 1966). With regard to the four types of hairs that characterize the hair coat, monotrichs, awls, auchenes, and zigzags are generated by hair follicles that are initiated in that order in groups (Claxton, 1966). Monotrichs come from the oldest follicles and zigzags from the youngest, with awls and auchenes in between (Claxton, 1966). The mechanism that regulates the production of the various hair types by follicles is largely unknown. It is known that several mutant genes interfere with the embryonic development of a hair follicle despite its initiation by cell signaling, while several other genes regulate the duration of the hair growth cycle in young and adult mice (Hebert et al., 1994; Lindner et al., 2000). Evidence that the embryology of hair growth is under tight genetic regulation is evident in the crinkled mouse. A recessive mutant allele at the crinkled (cr) locus produces a radical change in the hair coat of mice. Guard hairs, auchenes, and zigzags are missing from the thin hair coat of crinkled mice. The only hairs present are modiﬁed awls. Their hairs display many wrinkles absent in normal awls. Their hairs are thinner and the pigment deposits in their hair are more irregular than in normal awls. The agouti pattern of crinkled mice is altered. The dorsum tends to be darker and the ﬂanks yellower than in normal wild-type mice, apparently the result of alterations 81

CHAPTER 3

in the amount and distribution of black and yellow hair pigment as well as the disorderly appearance of the crinkled hair coat in general (Gruneberg, 1952). Straile and coworkers (Straile, 1969; Straile and Mann, 1973) have provided a model for the structural and functional organization of mammalian skin. He proposes that their skin is divided into numerous subunits, each containing nerves, blood vessels, hair follicles, and glands organized into a complex coordinated vertical system. A major component of each unit is a tylotrich hair follicle. Tylotrichs are atypical guard hairs/monotrichs. Tylotrich hair follicles are richly innervated, produce coarse hairs, and function as mechanoreceptors. They are thought to have other more complex neurophysiologic functions as well. Straile has noted that the richly innervated tylotrich follicles are typically surrounded by pigmented networks of melanocytes, suggesting that nerve ﬁbers servicing the hair follicle may stimulate melanocyte melanogenesis directly or lower the threshold for melanocyte melanogenesis in response to stimulation by endogenous or exogenous regulatory agents (e.g. hormones and growth factors). In contrast, dermal melanocyte networks associated with small, poorly innervated hair follicles are thought to be amelanotic because of the lack of neurological stimulation that also sets a higher threshold for their proliferating and “melaninizing” in response to intrinsic nonneural stimulation by the agents already mentioned. Only powerful exogenous factors, such as high doses of sex hormones or tumor promoters, can unleash these activities. For example, perifollicular melanocyte networks are more numerous in the trunk skin of female compared with male hamsters, indicating some response to estrogens (Ghadially and Illman, 1964; Illman and Ghadially, 1966). Consistent with Straile’s “threshold theory” is the fact that administration of estrogen to male hamsters increases the number of perifollicular melanocytes considerably beyond that of normal females of comparable age (Ghadially and Illman, 1964; Illman and Ghadially, 1966). Additional support for Straile’s threshold model for melanocyte activation and responsiveness to nonneural regulatory factors comes from the demonstration that the interruption of the adrenergic nerve supply to the irides of rabbits results in reduced tyrosinase activity within stromal melanocytes of the iris (Laties and Lerner, 1975). Within tylotrich units, the perifollicular melanocytes may inﬂuence other cellular components by secreting a variety of cytokines and matrix molecules (Chapter 21).

The Developmental Genetics and Regulation of Melanoblast Migration General Embryology: the Neural Crest Origin of Melanoblasts and their Migration to Populate the Skin of the Developing Wild-Type Mouse In mice, embryonic development is timed from copulation (coitus), indicated by the presence of a vaginal plug usually found in the vulva of females. It results from the coagulation of the male ejaculate. The timing of embryonic events below is expressed as the number of days post coitus (dpc). 82

During development, neural crest cells arise on the dorsal ridges of the neural folds that fuse to form the neural tube, precursor of the brain and spinal cord, etc. The neural crest cells give rise to many different types of cells, the melanocytes being one of them (Chapter 5). The precursors of the epidermal melanocytes are known as melanoblasts. They start to migrate dorsolaterally from the trunk region of the neural crest at 8.5 days, by which time the bilateral neural ridges of the embryo have already fused to form the neural tube (Serbedzija et al., 1990). The melanoblasts migrate in contact with the extracellular matrices surrounding the neural tube and “somites” and enter the mesenchymal layer (presumptive dermis) situated below the ectoderm (presumptive epidermis). Within the dermal mesenchyme, they migrate to all regions of the developing skin including that of the appendages (Pavan and Tilghman, 1994). By 10.5 days, neural crest cells are no longer seen leaving the neural crest (Serbedzija et al., 1990). By approximately 12 days, melanoblasts populate essentially all regions of the developing skin, including that of the limb buds (Mayer, 1973). With migration almost completed, the melanoblasts begin to populate the epidermis on day 12, where they proliferate and differentiate into melanocytes (Mayer, 1973). Apparently, many of the melanocytes migrate into the developing hair follicles. Those that do not remain in the basal layer of the epidermis. While the epidermal melanocytes establish intimate associations with keratinocytes of the interfollicular epidermis, the melanocytes of hair follicles do the same with respect to keratinocytes destined to form the hair produced by the follicle. By 16 days, melanocytes with dopa-positive premelanosomes are present within hair follicles (Hirobe, 1995). Throughout the migration of melanoblasts from the neural crest, there is no guarantee that melanoblasts will survive to complete their journey. Even melanocytes that enter the epidermis of the skin of mouse embryos are not immune to death. In the hairy skin of the trunk of mice, most of the interfollicular epidermal melanocytes that did not enter the hair follicles essentially disappear from the epidermis of the trunk skin by the 14th day post partum (Quevedo et al., 1966; Weiss and Zelickson, 1975). Electron microscopy indicates that the interfollicular epidermal melanocytes degenerate and die, questionably by apoptosis, a programmed event resulting in cell death (Quevedo et al., 1976; Weiss and Zelickson, 1975). However, the hair follicle melanocytes persist in the same areas where the epidermal melanocytes perish. In contrast to the trunk skin, the hair cover of the external ear of mice is sparse. The dark color of the outer ear results from melanin produced by epidermal and dermal melanocytes that persist throughout life.

Regulation of the Development of the Pigmentary System Embryogenesis is regulated by interactions at the subcellular, cellular, and tissue levels of organization. Growth factors,

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

signal transduction mechanisms, and transcription factors play essential roles orchestrating development. Substantial numbers of growth factors secreted by paracrine cells stimulate target cells to proliferate and often to differentiate. Signal transduction mechanisms acting within the target cell transform the stimulus provided by a growth factor into a new one capable of provoking an appropriate response by the target cell. Transcription factors interact with short sequences of chromosomal DNA situated in the enhancer and promoter regions of a gene in order to solicit its transcription into RNA and, indirectly, its translation into protein. Regulation of the distribution of melanoblasts from the neural crest of the mouse to the appropriate sites required to establish the pigmentary system begins with release of melanoblasts along with other neural crest cells from bondage in the neural crest. Initially, the neural crest cells are bound together by tight junctions that prevent their dispersal. In order to do so, these bonds have to be broken. This is brought about by the expression of BMP4 and BMP7 (bone morphogenetic protein) genes initiating a chain of events that includes the stimulation of the production of Slug gene and RhoB gene proteins. One of them activates factors capable of breaking the bonds of tight junctions (Slug), and the other (RhoB) brings about the enhancement of neural crest cell motility (RhoB). The removal of cell adhesion molecules from the surfaces of neural crest cells completes their preparation for invading regional environments on their way to those where they ﬁnally lodge. Depending on their gene expression, these environments may play an essential role in determining the fate of the melanoblasts (Gilbert, 2000; Markert and Silvers, 1956). The hair coat of the wild-type house mouse is pigmented throughout, indicating that the melanoblasts that migrate from the neural crest encounter hospitable environments during their journey. These supportive environments encountered by wild-type melanoblasts include regions of extracellular matrix lacking migration inhibitors but containing proteins such as proteoglycans capable of supporting melanoblast migration toward the presumptive skin. They encounter paracrine factors such as endothelins, notably endothelin-3, that direct the melanoblasts toward the skin and instruct them to differentiate as melanocytes there (Baynash et al., 1994; Chapter 13). On entering the skin, the melanoblasts encounter paracrine Wnt protein, which is secreted by mesenchymal cells and serves to reinforce the commitment of melanoblasts to become melanocytes (Dorsky et al., 1998). The substitution of some mutant alleles for their wild-type allele can drastically alter the generation of pigment in skin and hair follicles by blocking melanin synthesis. Substitution of other mutant alleles for their wild-type counterparts may produce less dramatic absence of melanin. Still other mutant alleles alter the color of melanosomes or their distribution in the hair. Collectively, the mutant alleles reveal the role that individual wildtype genes play in the formation of the pigmentary system and the complexity of their interaction. For example, homozygous Steel (Sl/Sl) mutants are charac-

terized by severe macrocytic anemia, rarely survive to birth, and are completely deﬁcient in germ cells as well as neural crest-derived melanocytes. The early death of Sl/Sl mice makes study of their deﬁciencies difﬁcult. However, the Dickie mutation (Sld) of the Steel locus, in combination with the Sl mutant allele, yields a Sl/SId mouse that is sterile, anemic, and reduced in life expectancy (may survive up to a year) and, except for its black eyes, unpigmented. Melanocytes are missing from their skin and hair follicles (Silvers, 1979). Such mice are capable of living long enough for study of the consequences of stem cell deﬁciencies resulting from mutations at the Steel locus. Fujita et al. (1989) have demonstrated that the mutant Sl/Sld ﬁbroblasts fail to produce and secrete a growth factor, Steel factor, better known as stem cell factor (SCF), essential for the proliferation and differentiation of stem cells, which give rise to reproductive cells, blood cells, and melanocytes. It is also important in regulating the proliferation and differentiation of neonatal mouse epidermal melanocytes in culture (Hirobe et al., 2004). In wild-type mice, SCF is active and binds to a transcell membrane melanoblast receptor, c-kit tyrosine receptor kinase (c-kitTRK), a product of the Dominant spotting W locus. The binding of SCF protein (ligand) to its receptor c-kitTRK activates the receptor, resulting in the onset of a sequence of reactions that activates the Mitf transcription factor encoded by the microphthalmia gene. As a result, Mitf transcription factor combines with a protein coactivator. The combination of activated Mitf and coactivator is capable of activating the transcriptions of genes central to the production of melanin (Price et al., 1998). In contrast to wild-type SCF of ﬁbroblast origin that exerts an environmental inﬂuence on wild-type melanoblasts, the Dominant spotting W locus is expressed within the melanoblast (Silvers, 1979). Like the Steel mutant, Dominant spotting (W/Wv) mice have black eyes and are otherwise unpigmented, sterile, and anemic. The defective proliferation, differentiation, and survival of the W/Wv melanoblasts results from their failure to respond to appropriate cues from their tissue environment, namely by binding and responding to potent SCF. The fault resides in their defective c-kit tyrosine receptor kinase that fails to respond to SCF. As a result, c-kitTRK does not autophosphorylate and activate functions essential for normal development (Nocka et al., 1989). Injection of anti-c-kitTRK antibody (ACK2) into pregnant black (C57BL/6) mice reveals a c-kit-dependent stage of melanoblast development on day 14.5 of gestation, just before c-kitTRK-dependent melanoblasts enter the epidermis from the dermis (Nishikawa et al., 1991). In addition, treatment of adult black mice with ACK2 indicates a requirement for ckitTRK action on follicular melanocytes at the onset of each hair growth cycle. It seems likely that c-kitTRK activity is required to promote melanoblast proliferation within the embryonic dermis and melanocyte proliferation at the onset of each hair growth cycle (Nishikawa et al., 1991). Okura et al. (1995) found that c-kitTRK dependency was not restricted to a brief period in the dermis prior to melanoblast invasion of 83

CHAPTER 3

the epidermis, but persisted during their differentiation into melanocytes and migration into the hair follicles. As noted above, the absence of neural crest-derived melanocytes in the skin and hair of Steel mutant (Sl/Sld) mice and Dominant spotting (W/Wv) mice results from the failure of ﬁbroblasts of Steel mutant mice to deliver SCF. Alternatively, as in the case of mutant Dominant spotting mice, the ﬁbroblasts’ SCF signal is available to melanoblasts, but they lack the competent c-kit tyrosine receptor kinase required to register and react to it. The impact of mutations at the Steel and Dominant spotting loci on the development of the neural crest-derived melanin pigmentary system has been examined. The goal was not only to determine what went wrong but also to obtain hints as to the role that each wild-type gene plays in establishing the wild-type pigmentary phenotype. A major problem that investigators have encountered when tracing the migration of melanoblasts from the neural crest is the close resemblance between melanoblasts and the population of mesenchymal cells through which they migrate on their way to the epidermis and hair follicles of the skin. A particularly useful method for making this distinction is the construction of probes that bind to speciﬁc molecules, often proteins that are unique to melanoblasts and melanocytes. One such protein is tyrosinase-related protein #2 (TYRP-2), an enzyme that plays an important role in the synthesis of melanin (Pavan and Tilghman, 1994; Steel et al., 1992). It is produced relatively early in the migration of melanoblasts even though melanogenesis is a late event that characterizes the transformation of melanoblasts into melanocytes. TYRP2 protein is associated with the limiting membrane of melanosomes, the cell organelles within which melanogenesis takes place. Please remember that, in the research described below, wherever melanoblasts and melanocytes are mentioned, they were judged to be so based on a positive response to a TYRP-2 protein probe, tyrosinase mRNA probe, TYRP1 mRNA probe, or TYRP-2 mRNA probe. With regard to the time of ﬁrst detection of neural crest melanoblasts, TYRP-2 was ﬁrst with day 10 dpc, followed by TYRP-1 (day 14.5 dpc), tyrosinase (day 14.5 dpc), and pigment deposition (day 16.5 dpc) (Pavan and Tilghman, 1994; Steel et al., 1992). Steel et al. (1992) emphasized the TYRP-2 mRNA probe in their study because of its early detection of neural crestderived melanoblasts and its ability to distinguish between migrating melanoblasts and mesenchymal cells in their study of Steel mice. They found that SCF characteristic of wild type was absent in Sl/Sld mutant mice, a deﬁcit that led to premature melanoblast/melanocyte death or dedifferentiation beginning around day 11 dpc and continuing thereafter, resulting in a uniformly white coat of hair. Predictably, in mice homozygous for the wild-type allele, the hair coat was pigmented throughout. The study also revealed that, although SCF signaling is essential for the survival of melanoblasts, it does not play a role in their early migration or initial differentiation. Steel et al. (1992) (see also their communication to Pavan and Tilghman published in their 1994 paper) explored melanoblast fates in mutant mice of the Dominant spotting W 84

locus. The genotypes examined were Wv/Wv, Wsh/Wsh, and W41/W41. All were characterized by white hair coats due to melanoblast/melanocyte death by day 12 dpc, a consequence of their lack of a c-kit tyrosinase receptor kinase responsive to SCF signaling. The study also revealed that, although c-kit tyrosinase receptor kinase is essential for the survival of melanoblasts, it does not play a role in their early migration or initial differentiation. Pavan and Tilghman (1994) built on the foundation laid by Steel et al. (1992) when they used an antibody speciﬁc for the TYRP-2 protein as a probe to identify the function of the piebald s locus by examining the inﬂuence of the piebald lethal mutant allele (sl) on the fate of neural crest-derived melanoblasts. In wild-type mice by day 12 dpc, numerous melanoblasts/melanocytes are localized in the head region back to the forelimbs followed by numerous melanoblasts from the hindlimbs back to the tail. The region between the limbs, the trunk of the body, has signiﬁcantly fewer melanoblasts than the regions fore and aft until days 13–17 dpc when a wave of mitotic activity sweeps across the trunk increasing melanoblast numbers to levels comparable to those elsewhere along the anterior–posterior axis. This is not the case in piebald lethal sl/sl mice. Typically, variable patches of pigmented hair are found on the head, rump, or base of tail, with the space between them populated by white hair. From day 10.5 dpc, when neural crest-derived melanoblasts are seen, their numbers are extremely low compared with wild type. In piebald lethal mice, melanoblasts were only found in regions of the skin where patches of pigmented hair were frequently observed. As the numbers of melanoblasts found in the skin of piebald lethal mice was low from their ﬁrst appearance on day 10 dpc, Pavan and Tilghman (1994) concluded that the s locus wild-type gene must express early in melanocyte development either in the neural crest or during melanoblast migration. They suggest that the expression of the piebald locus is upstream from the Steel and Dominant spotting loci. In an effort to integrate the interaction of Steel, Dominant spotting, and Microphthalmia signals, a provisional pathway for the developmental genetic events that culminate in the onset of melanogenesis within differentiating melanoblasts has been assembled: ﬁbroblasts homozygous for the wild-type Steel gene that encodes a protein, SCF, secrete it into the dermis during embryogenesis. Its target is the melanoblasts arriving from the neural crest. SCF becomes bound to the melanoblast cell surface c-kit receptor tyrosine kinase (ckitRTK) encoded by the wild-type allele of the Dominant spotting W locus. The binding of SCF to the c-kitRTK dimerizes the receptor, leading to its phosphorylation. This event is critical in the activation of a pathway resulting in the phosphorylation of the wild-type microphthalmia locus transcription factor (Mitf). The phosphorylated form of Mitf is capable of initiating a reaction that results in the onset of transcription and translation of the wild-type alleles for the enzymes essential in the synthesis of melanin: tyrosinase, DHICA oxidase/TYRP-1, and DOPAchrome tautomerase/TYRP-2, and other gene products relevant to melanogenesis.

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

It is reasonable to conclude that a major function of the wild-type alleles at many of the mutant “white-spotting” gene loci, including the Dominant spotting locus, is to orchestrate melanoblast migration, proliferation, and differentiation via inﬂuences on the availability of growth factors and by determining the response of melanoblasts to them. The physiological genetics of the Steel locus and Dominant spotting W locus stand as a classic model for such interactions.

The Regulation of the Fully Differentiated Melanocyte in Producing the Wild-type Phenotype The production of hairs with the appropriate pigmentation to produce the agouti pattern requires close timing of the growth of the hair and the function of melanocytes that pigment it. Although some of the regulatory mechanisms involved have been identiﬁed, much remains uncertain. In wild-type agouti mice, the melanocytes of hair follicles programmed to produce banded hairs shift their synthesis of pigment from eumelanin to pheomelanin and back to eumelanin during hair growth. As a result, melanin is deposited in viable keratinocytes destined to keratinize and die while forming the hair shaft. In doing so, they also take part in coloring the hair. The regulation of melanocyte function in this process is as follows. The signal for the production of pheomelanin is provided by agouti protein encoded by the wild-type allele of the agouti a locus activated within dermal cells that form the dermal papilla of hair follicles (Jackson, 1994; Chapter 19). Within the hair bulb, melanocytes are arranged so that they are adjacent to the dermal papilla on one side and to keratinocytes on the other. Alpha-melanocyte stimulating hormone (MSH), also known as melanocortin I, binds to a specialized cell surface receptor on the melanocytes, the MSH receptor encoded by the wild-type allele of the extension locus (e) (Jackson, 1994). The receptor-bound MSH stimulates events that result in the activation of adenyl cyclase, which elevates the levels of cyclic adenosine monophosphate (AMP), leading to the synthesis of eumelanin. The eumelanin-containing melanosomes are transferred to the population of keratinocytes destined to form the hair tip (Boissy, 2003; Marks and Seabra, 2001). Subsequently, agouti protein emanating from the dermal papilla cells gains access to the MSH receptor on the melanocytes and blocks the binding of MSH to them. Eumelanin synthesis is halted and pheomelanin synthesis is commenced within melanocytes. The pheomelanosomes are transferred to the population of keratinocytes scheduled to form the agouti band. Finally, MSH replaces agouti protein on the receptors, and eumelanin is synthesized and distributed to keratinocytes scheduled to form the remainder of the hair shaft. Hair growth stops and developmental changes occur so that the hair is elevated and anchored in the now quiescent follicle. While this description accurately relates the expression of the wild-type agouti gene, some agouti alleles regulate the dorsum and

ventrum of the mouse independently, yielding different color patterns than wild type (Millar et al., 1995; Vrieling et al., 1994). Within the follicular melanocytes during the periods of eumelanin synthesis in agouti mice, numerous genes are activated to support the process. The synthesis of eumelanin involves a pathway whereby tyrosine is converted via intermediates into the complex polymer eumelanin. Not surprisingly, the transcription of genes that encode enzymes that catalyze the melanogenic pathway is activated: the wild-type allele at the albino locus (c) encoding tyrosinase, the wild-type allele at the brown locus (b) encoding the protein TYRP-1 (DHICA oxidase), and the wild-type Slaty locus (Slt) allele encoding TYRP-2 (DOPAchrome tautomerase) (Chapter 10). In addition, the wild-type allele at the pink-eyed dilution locus (p), a gene that encodes a protein that is not an enzyme, is also activated. It is believed that all the foregoing proteins are aggregated to form a “melanogenic complex” that is bound to the limiting membrane of the melanosome. The “melanogenic complex” is postulated to generate the eumelanin that is deposited on and between the protein ﬁlaments of a well-organized matrix that occupies the interior of the eumelanosome. Possible functions of the wild-type p protein might be that it functions as a structural element of the melanosome, a coordinating molecule of the “melanogenic complex,” a transporter of melanogenic proteins, or is involved in the transport of sulfhydryl compounds out of the melanosome to permit the formation of eumelanin (Lamoreux et al., 1995). During the pheomelanic phase of melanogenesis in the agouti mouse brought on by binding of agouti protein to the MSH receptors on follicular melanocytes, the biochemistry of melanin formation takes a new direction. Pheomelanin synthesis begins with the oxidation of tyrosine catalyzed by tyrosinase, but then diverges to interact with sulfur-containing compounds such as glutathione or cysteine to form the sulfurous compound of cysteinylDOPA. CysteinylDOPA is modiﬁed and then polymerized to form pheomelanin (Chapter 10). These events take place within pheomelanosomes that lack the more complex internal organization found in eumelanosomes. Pheomelanosomes are spherical in shape, whereas eumelanosomes are ellipsoidal. The “melanogenic complex” of eumelanosomes is absent in pheomelanosomes with only tyrosinase activity in evidence in the four members of the “melanogenic complex.” Microvesicles dominate the interior of the pheomelanosome, and protein ﬁlaments, where present, are scattered rather than arrayed to form an organized matrix. Pheomelanin is deposited on the surfaces of the microvesicles and ﬁlaments and ﬁlls the space between them. Tyrosinase may be present within the microvesicles where it converts tyrosine to dopaquinone, which diffuses out of the vesicles and interacts with cysteine or glutathione to form cysteinylDOPA that, in modiﬁed form, is polymerized into pheomelanin. The pheomelanosomes are discharged into the keratinocytes scheduled to form the agouti band (Chapter 19; Jimbow et al., 1999). 85

CHAPTER 3

Examples of Inﬂuences of Hair Follicle Growth Regulators on the Hair Growth Cycle with Possible Implications for Follicular Melanocytes Fibroblast growth factors (FGFs) form a family, the members of which have been described as possibly the most important “endogenous mediators of hair follicle growth and development” (Danilenko et al., 1995). The family includes acidic FGF (aFGF), basic FGF (bFGF), and FGF-5, the receptors for which are found within hair follicles or very close to them. Another member of the family (KGF or FGF-7) and its receptor, although localized to the skin, inﬂuences hair growth (Danilenko et al., 1995). Halaban in Chapter 22 and Hirobe (1992) have also stressed the importance of the members of the FGF family for their role as supporters of melanocyte proliferation and viability in vitro. Halaban notes that, although bFGF is the most potent supporter of melanocyte proliferation/survival in vitro, other members can substitute for it. The question that remains to be answered is whether the commitment of FGFs to regulating hair growth permits their simultaneously supporting follicular melanocytes of the hair bulb that pigment the developing hair shaft while performing largely unknown functions in the ORS. One case study says no, not because the growth factor is necessarily impotent but because, in meeting the hair growth cycle’s requirement, it is expressed too late, that is, just as hair growth is about to stop. Fibroblast growth factor-5 (FGF-5) expresses and is sharply localized in the external root sheaths of hair follicles late in anagen (Hebert et al., 1994). Evidence suggests that FGF-5 is an essential component of the signaling system that elicits the end of anagen and onset of catagen and terminates each hair growth cycle. Supporting this role for FGF-5 is the demonstration that genetically engineered mice homozygous for a null (nonfunctional) allele of FGF-5 (FGF5neo) produced by gene targeting in embryonic stem cells have longer hairs than unaltered mice. In addition, Angora (go) mice are indistinguishable from FGF5neo mice in their coat of elongated hair, compared with that of wild-type mice. The Angora gene is considered to be an allelic mutant of the FGF-5 gene (Hebert et al., 1994). The late expression of the FGF-5 gene and its restriction to the ORS argues against any inﬂuence on the hair bulb melanocytes, nor has there been a report of its inﬂuence on melanocytes of the ORS. Examination of the timing of gene expression and distribution patterns of other FGFs may reveal one or more that meets the needs of melanocytes and the growth of the hair. Another candidate for a growth factor inﬂuencing both hair growth and melanocytes is hepatocyte growth factor (HGF) and its receptor MET. The team is known to be an important mediator of mesenchymal–epithelial interactions and involved in the control of hair growth (Lindner et al., 2000). HGF is restricted to the dermal papilla of hair follicles, and its receptor MET is present nearby in keratinocytes of the hair bulb. 86

HGF has been shown to repress the onset of the catagen stage of the hair growth cycle, thus prolonging it (Jindo et al., 1994; Lindner et al., 2000). Consequently, HGF action on hair growth is just the opposite of that of FGF-5. While HGF acts to delay the onset of catagen, FGF-5 acts to promote its onset. In this case, the duration of hair growth might be a compromise between the antagonistic inﬂuences. HGF is known to act either alone (Hirobe et al., 2004) or synergistically with bFGF in supporting melanocyte proliferation in vitro (Chapter 22). The distribution pattern of HGF and its receptor in growing hair follicles, as well as its expression beginning early in anagen, are consistent with a possible role in supporting the proliferation and viability of follicular melanocytes during the hair growth cycle, particularly those of the hair bulb.

Conclusion The wild-type agouti coat color is but one pattern of the collective expression of gene loci involved in pigmenting the skin and hair. Several of the chapters that follow this one explore in greater depth the mutant alleles at these loci that, when present, result in departures from the wild-type color pattern. In doing so, they help to reveal the nature of the steps in the pigmentary process that a particular gene locus inﬂuences. Signiﬁcant numbers of mutant “pigment genes” have been characterized, not only for their gross inﬂuences on coat color but also for their actions at the molecular level. Only a sample has been discussed in this chapter. The same is true for the hormones, growth factors, signal transduction mechanisms, and other factors that regulate melanin synthesis by melanocytes. More information on these and other topics can be found in the chapters listed below. Human Skin Color Chapter 20: Human Pigmentation: Its Regulation by Ultraviolet Light and by Endocrine, Paracrine, and Autocrine Factors Chapter 27: The Normal Color of Human Skin Melanin and Human Adaptations to Solar Radiation Chapter 17: Photobiology of Melanins The Embryogenesis of the Melanin Pigmentary System Chapter 5: Regulation of Melanoblast Migration and Differentiation Chapter 6: Melanoblast Development and Associated Disorders Chapter 7: Biogenesis of Melanosomes Genetics of the Pigmentary System Chapter 10: The Regulation of Melanin Formation Chapter 11: The Tyrosinase Gene Family Chapter 19: Regulation of Pigment Type Switching by Agouti, Melanocortin Signaling, Attractin, and Mahoganoid

GENERAL BIOLOGY OF MAMMALIAN PIGMENTATION

The Regulation of the Fully Differentiated Melanocyte Chapter 10: The Regulation of Melanin Formation Chapter 11: The Tyrosinase Gene Family Chapter 12: Molecular Regulation of Melanin Formation: Melanosome Transporter Proteins Chapter 14: Enzymology of Melanin Formation Chapter 15: Chemistry of Melanins Chapter 20: Human Pigmentation: Its Regulation by Ultraviolet Light and by Endocrine, Paracrine, and Autocrine Factors Chapter 22: Growth Factor Receptors and Signal Transduction Regulating the Proliferation and Differentiation of Melanocytes

Perspectives This chapter has surveyed the products of Jacob’s (1982) evolutionary tinkering that transformed the pigmentary system of reptiles into that of mammals. The great diversity of coat color patterns found in mammals represents developmentally subtle manipulations of a pigmentary system that appears to be highly conserved at its core. The basic ingredients for hair and skin color (melanocytes, melanosomes, melanogenic enzymes, and melanin) seem to be pretty much the same throughout the class of mammals, as does the basic ability of melanocytes to transfer melanosomes to keratinocytes, an attribute inherited from the reptilian ancestors of mammals. What is different are the regulatory factors that program the timing of pigment pattern expression, the anatomical locations of hair follicles and the coarseness of the hairs that are produced, and the qualitative and quantitative functions of melanocytes. More is being learned daily about these regulatory systems. The importance of the melanin pigmentary system of mammals cannot be overemphasized, for it is at the interface between the organism and the world in which it must survive and reproduce. What mammals have inside them is insigniﬁcant unless it is in harmony with their integument. Mammals communicate their strengths and desires through their color patterns as well as concealing their vulnerability. In the animal kingdom, color patterns form a powerful silent language focused on aggression, attraction, repulsion, submission, and aloofness. Tragically, with all its capacities for doing good things, taken as a whole, the human species has still not been able to penetrate beneath the superﬁciality of skin color in assessing the true worth of its individual members.

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Silvers, W. K. The Coat Colors of Mice. A Model for Mammalian Gene Action and Interaction. New York: Springer-Verlag, 1979. Steel, K. P., D. R. Davidson, and I. J. Jackson. TRP-2/DT, a new early melanoblast marker, shows that steel growth factor (c-kit ligand) is a survival factor. Development 115: 111–119, 1992. Straile, W. E. Carcinogen-induced melanotic tumours of the tylotrich hair follicle. Nature 202:403–404, 1964. Straile, W. E. Vertical cutaneous organization. J. Theor. Biol. 24: 203–215, 1969. Straile, W. E., and S. J. Mann. Characteristics of hairs and related mechanoreceptors in the mouse and rabbit. J. Exp. Zool. 185:189– 208, 1973. Sugiyama, S., T. Uesugi, K. Jimbow, and A. Kukita. Activation and differentiation of melanogenesis in melanocytes of telogen and early anagen hair follicles. In: Pigment Cell, vol. 4, S. Klaus (ed.). Basel: S. Karger, 1979, pp. 185–190. Szabo, G. The regional anatomy of the human integument with special reference to the distribution of hair follicles, sweat glands and melanocytes. Philos. Trans. Roy. Soc. London 252:447–485, 1967. Szabo, G., A. B. Gerald, M. A. Pathak, and T. B. Fitzpatrick. The ultrastructure of racial color differences in man. In: Pigmentation: Its Genesis and Biologic Control, V. Riley (ed.). New York: Appleton Century Crofts, 1972, pp. 23–41. Thody, A. J., E. H. Higgins, K. Wakamatsu, S. A. Burchill, and J. M. Marks. Pheomelanin as well as eumelanin is present in human epidermis. J. Invest. Dermatol. 97:340–344, 1991. Thong, H.-Y., S.-H. Jee, C.-C. See, and R. E. Boissy. The pattern and mechanism of melanosome distribution in keratinocytes of human skin is one determining factor of skin colour. Br. J. Dermatol. 149:498–505, 2003. Vrieling, H., D. M. J. Duhl, S. E. Millar, K. A. Miller, and G. S. Barsh. Differences in dorsal and ventral pigmentation result from regional expression of the mouse agouti gene. Proc. Natl. Acad. Sci. USA 91:5667–5671, 1994. Waage, J. K. How the zebra got its stripes — Biting ﬂies as selective agents in the evolution of zebra coloration. J. Entomol. Soc. S. Afr. 44:351–358, 1981. Weiss, L. W., and A. S. Zelickson. Embryology of the epidermis. III. Maturation and primary appearance of dendritic cells in the mouse with mammalian comparisons. Acta Dermatovener 55:431–442, 1975. Whimster, I. W. An experimental approach to the problem of spottiness. Br. J. Dermatol. 77:397, 1965. Whimster, I. W. Symmetry in dermatology. In: Modern Trends in Dermatology — 4, P. Borrie (ed.). London: Butterworths, 1971, pp. 1–31. Wickings, E. J., and A. F. Dixson. Testicular function, secondary sexual development and social status in male mandrills (Mandrillus sphinx). Physiol. Behav. 52:909–916, 1992. Wickler, W. Socio-sexual signals and their intra-speciﬁc imitation among primates. In: Primate Ethology, D. Morris (ed.). London: Weidenfeld and Nicolson, 1967, pp. 69–147. Witkop, C. J., Jr., W. C. Quevedo, Jr., T. B. Fitzpatrick, and R. A. King. Albinism. In: The Metabolic Basis of Inherited Disease, 6th edn, vol. II, C. R. Scriver, A. L. Beaudet, W. S. Sly, and D. Valle (eds). New York: McGraw-Hill, 1989, pp. 2905–2947. Yamamoto, O., and J. Bhawan. Three modes of melanosome transfers in Caucasian facial skin: hypothesis based on an ultrastructural study. Pigment Cell Res. 7:158–169, 1994. Yang, J.-S., R. M. Lavker, and T. T. Sun. Upper human hair follicle contains a subpopulation of keratinocytes with superior in vitro proliferative potential. J. Invest. Dermatol. 101:652–659, 1993. Young, A. R. The sunburn cell. Photodermatology 4:127–134, 1987. Yun, J. S., and W. Montagna. The melanocytes in the epidermis of the rhesus monkey (Macaca maculatta). Anat. Rec. 154:161–174, 1966. Zuckerman, S., and A. S. Parkes. Observations on secondary sexual characters in monkeys. J. Endocrinol. 1:430–439, 1939.

The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

4

Extracutaneous Melanocytes Raymond E. Boissy and Thomas J. Hornyak

Historical Background Our analysis and knowledge pertaining to the structure/ function correlates of pigment and melanocytes/chromatophores have primarily focused on the various populations of pigment-synthesizing cells in cutaneous tissues. These populations of cells have included the dermal melanophores of lower vertebrates, which exhibit rapid melanosome dispersion or constriction within the cell as an immediate adaptation to the environment. In addition, other chromatophores of lower vertebrates, in association with the melanocyte, interact in the dermis and at the dermal/epidermal junction to provide an array of cutaneous coloration for environmental and social interactions. Follicular melanocytes in the bulb region of the hair follicles of higher vertebrates and primates are responsible for making melanosomes that are transferred to the hair keratinocytes and thus give coloration to the hair shaft, which presumably functions to provide cues to and possibly protection from the environment. Interfollicular epidermal melanocytes are primarily, but not exclusively, found in most primates including humans. This population of melanocytes developed in conjunction with the dramatic decrease in the terminal type of hair follicles distributed throughout the body surface. These interfollicular melanocytes of the epidermis seem to have evolved for additional and/or different functions, including the absorption of ultraviolet and visible light and the scavenging of free radicals. Melanin, eventually transferred to keratinocytes, appears to protect the body in general, and the skin speciﬁcally, against damaging environmental assaults. Thus, looking globally at the evolutionary development in the various populations of cutaneous pigment cells, one can see that the functions of melanin change or increase as described above. However, all these various functional roles for cutaneous melanocytes reﬂect mechanisms responsible for the interaction between an individual and the environment. Less has been known about extracutaneous melanocytes found primarily in the eye, ear, and internal organs such as the leptomeninges. These populations of melanocytes have some morphologic and functional similarities to the various cutaneous melanocytes outlined above. However, there are some signiﬁcant differences between extracutaneous and cutaneous melanocytes that have yet to be fully assessed. It appears that extracutaneous melanocytes must have unique functional roles for the organism distinct from those of cutaneous melanocytes by virtue of the fact that extracutaneous melanocytes are not positioned immediately at the organ-

ism/environmental interface. This precludes the function of extracutaneous melanocytes as primarily providing environmental protection and/or socially interactive cues. A review of the characteristics of extracutaneous melanocytes is presented in this chapter.

Development and Nature of Ocular Melanocytes Ocular melanocytes have traditionally been categorized into two groups of cell types: the uveal melanocytes and the retinal pigmented epithelial (RPE) cells. It should be mentioned that a controversy exists as to whether the RPE cells are truly melanocytes. It has been proposed that, as these cells express melanocyte-speciﬁc proteins for the specialized synthesis of pigment granules and melanin, they are by deﬁnition “melanocytes.” Conversely, it has been argued that these cells that form a continuous epithelial tissue in association with the retina proper, thus forming a border or lining between the body (i.e. mesenchyme tissue of the uvea) and a body cavity (i.e. vitreous chamber), are actually epithelial cells. This proposal presumes that the synthesis of melanin by these epithelial cells is an accessory function. The reality is probably somewhere in between, as the epithelial function of the RPE is important for eye development whereas its role in pigment synthesis is critical for proper retinal ganglion cell pathﬁnding. We will refer to these cells as RPE melanocytes. In any case, the uveal and RPE categories of ocular melanocytes can be distinguished from each other on both a developmental and a morphologic basis. For comprehensive reviews of ocular embryology and structure, we recommend Jakobiec (1982) and Fine and Yanoff (1979). Developmentally, melanocytes that eventually reside in the uvea, comprising the choroid, ciliary bodies, and the iris, are of neural crest origin similar to cutaneous melanocytes (see Chapter 5). Neural crest-derived melanoblasts migrate through the mesenchyme of the developing embryo and are subsequently targeted to this ocular site (Bartelmex and Blount, 1954; LeLievre and LeDouarin, 1975; Noden, 1991). In contrast, cells of the RPE are derived directly from cells in the neuroectoderm of the developing forebrain of the embryo (Mann, 1964). Speciﬁcally, after the diencephalon (forebrain) forms the optic cup that invaginates into a bilayered structure, the outermost layer adjacent to the mesenchyme of the developing uveal tract differentiates into the RPE. Therefore, no migratory phase occurs for these RPE melanocytes. Again, 91

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it has traditionally been stated that these two groups of melanocytes originate from different embryonic sources. This is misleading in that the precursors for both groups (the neural crest cells and the posterior surface of the optic neuroectoderm) actually originate from the same source, i.e. the neural tube. Neural crest cells emigrate from the dorsal surface of the neural tube shortly after the neural fold closes (Le Douarin, 1982). The neural ectoderm of the developing forebrain, which gives rise to the RPE, is the cephalic extension of the neural tube. Therefore, the developmental divergence that separates these two groups of ocular melanocytes is not their original embryonic origin per se, but rather their mode of reaching the target site. Melanoblasts that differentiate into uveal melanocytes undergo a migratory phase from neural tube to target site; in contrast, melanoblasts that differentiate into RPE melanocytes do not undergo a migratory stage but evaginate from a speciﬁc region of the neural tube. Analysis of murine coat color mutants has provided important insights into how neural crest-derived and RPE melanocytes are differentially regulated during development. The determinants of neural crest-derived melanocyte development are discussed elsewhere (Chapter 5) and should apply to the control of survival, migration, and initial differentiation of uveal melanocytes because of the evidence supporting their origin from the neural crest. Cell surface receptors such as Kit and Ednrb, their respective ligands KitL/SCF and ET-3, as well as the transcription factors Mitf, Pax3, Sox10, and Snai2 have all been shown to be important in governing the development of neural crest-derived melanocytes during embryogenesis. There are distinct differences between some of these determinants and determinants of RPE development. One example is the difference in Mitf isoform expression between neural crestderived melanocytes and RPE cells. Although Mitf is critical to the development of both types of cells, neural crest-derived melanocytes express the Mitf-M isoform exclusively, whereas RPE cells express the Mitf-A, Mitf-H, and Mitf-D isoforms, all expressed from different promoters and utilizing different ﬁrst exons (Takeda et al., 2002; Udono et al., 2000). As these isoforms are regulated differently, it follows that the transcriptional determinants for Mitf expression in RPE melanocytes are different than those for neural crest-derived melanocytes. In fact, neither Pax3 or Sox10, which regulate the expression of the neural crest-derived melanocyte isoform Mitf-M (Bondurand et al., 2000; Potterf et al., 2000), are expressed in RPE melanocytes. It appears that the major regulators of RPE melanocytes are the factors Mitf, Otx2, and Pax6 linked in a transcriptional network (Martinez-Morales et al., 2004), perhaps regulated by the activity of ﬁbroblast growth factors expressed by the overlying endoderm (Nguyen and Arnheiter, 2000). Additionally, mutations affecting the Kit or Ednrb systems inhibit successful migration of melanoblasts into the skin, the ear, and uveal tract without affecting survival or differentiation of RPE melanocytes (Cable et al., 1994; Deol, 1970a; Mackenzie et al., 1997; Markert and Silvers, 1956).

92

It was originally demonstrated in classic studies by Noden (1975) that the uveal melanocytes of the eye originate from neural crest cells derived from the cephalic portion of the neural tube. Two basic possibilities may explain the distinct differentiated phenotype of uveal melanocytes from cutaneous melanocytes. One possibility is that uveal melanocytes, although neural crest derived, are speciﬁed different from cutaneous neural crest-derived melanocytes and adopt their differentiated phenotype regardless of local environmental cues. The other possibility is that their speciﬁed precursor melanoblast is indistinguishable from those contributing to cutaneous melanocytes and that end-organ environmental signals are critical for conferring the differentiated phenotype. Some evidence exists to support each of these models. For example, uveal-like melanocytes have been demonstrated to emigrate from all areas of the neural tube (Boissy et al., 1990). A unique mutant allele at the albino (c) locus in Recessive white chickens affects the function of tyrosinase in cutaneous and not in uveal melanocytes (Brumbaugh et al., 1983; Oetting et al., 1985). Neural tubes extracted from 3-day-old embryos of this mutant line, trisected into cephalic, medial, and caudal segments, and used to develop separated cultures of melanocytes demonstrated that equal numbers of amelanotic and pigmented melanocytes could be derived from all areas of the neural tube. Therefore, the commitment to differentiate into cutaneous vs. extracutaneous melanocytes may be determined at some preneural crest cell stage. It has also been demonstrated that the normal embryonic development of the RPE and choroidal melanocytes is essential for the developmental establishment of the neural retina and its projection and connection to the visual cortex of the brain. Classic studies in all forms of albinism demonstrate that the associated innate hypopigmentation of the uveal tract and the RPE correlates with a signiﬁcant decrease in visual acuity associated with strabismus and nystagmus of the eyes (Creel et al., 1990; Fulton et al., 1978; King et al., 1994; Kinnear et al., 1985; Summers et al., 1988). These ocular/vision aberrancies are the consequence of a misrouting of the optic ﬁbers from the neural retina. There is an increase in neurons crossing at the optic chiasma to the contralateral geniculate, resulting in the scrambling of the neuronal input to the visual cortex of the brain (Creel et al., 1978; Guillery, 1974). Most intriguing is the fact that this misrouting of the visual system occurs in patients with OCA1, OCA2, or OCA3. These three forms of albinism result from nucleotide lesions in the gene encoding the tyrosinase enzyme (Chintamaneni et al., 1991; Giebel et al., 1990; King et al., 1991; Tomita et al., 1989), the P protein (Gardner et al., 1992; Ramsay et al., 1992; Rinchik et al., 1993), and tyrosinase related protein-1 (TRP-1) (Boissy et al., 1996) respectively. These three melanocyte-speciﬁc proteins regulate pigment synthesis and do not appear to directly affect any developmental process. Therefore, the aberrant visual system occurring in albinism demonstrates that the development of ocular melanin during embryogenesis is critical for, and probably directs, the normal establishment of the

EXTRACUTANEOUS MELANOCYTES

optic neuronal network (Colello and Jeffery, 1991; Silver and Sapiro, 1981; Strongin and Guillery, 1981) as well as photoreceptor cells directly, speciﬁcally rods (Grant et al., 2001). In mice, the number of rod photoreceptor cells that are produced decreases correlatively with extent of pelage hypopigmentation (Donatien and Jeffery, 2002). The mechanism underlying the regulation of the development of both the retina and the visual pathway by ocular melanins is still elusive (Ilia and Jeffery, 2000). Not only is the synthesis of melanin key in the development of the visual system of an organism, but the presence of RPE melanocytes is also responsible for the maintenance as well as the successful embryonic development and organization of the neural retina proper. Experimentally induced (Raymond and Jackson, 1995) or natural mutant (Boissy et al., 1993; Mullen and LaVail, 1976) perturbations in the development of the RPE during embryogenesis have correlated with the aberrant development of photoreceptor cells and other layers of the neural retina. Therefore, melanocytes and pigment development in the eye provide important cues for the development of the ocular and visual system. In addition, cells of the pigmented ciliary margin in mice have been shown to function as putative retinal stem cells (Tropepe et al., 2000), ascribing a potential postnatal role in retinal maintenance to a small subset of ocular pigment cells.

Morphology of Uveal (i.e. Choroidal) and RPE Melanocytes Once the uveal group and RPE group of melanocytes migrate to and/or differentiate in their respective anatomical sites, distinct morphologic criteria distinguish the two. These

differences are reﬂected in both the shape of the melanocytes (Fig. 4.1) and the morphology of the melanosomes synthesized. Pertaining to the cellular shape, the melanocytes of the uveal tract are dispersed throughout the connective tissue stroma of these areas in a territorial manner without any direct contact with each other (Feeney and Hogan, 1961). These uveal melanocytes are dendritic. They have a centrally placed nucleus in the cell body and peripherally radiating dendrites. Wholemounts of the choroidal component of the uveal tract have demonstrated that a majority of these melanocytes are bipolar; however, some uveal melanocytes develop numerous dendrites (Torczynski, 1982). The choroid is a layer of connective tissue per se of about 80 mm in thickness. The cellular component of the choroid is primarily composed of ﬁbroblasts and melanocytes in a 2:3 ratio and numerous endothelial cells which line the cavernous blood sinus of this optic layer (Fig. 4.2). Occasionally, nerve bundles containing groups of myelinated and nonmyelinated neurons exist (Fig. 4.2) (Bergmanson, 1977; Castro-Correia, 1967). These cellular components are embedded in a ﬁbrous stroma composed primarily of collagen and elastin ﬁbers (Feeney and Hogan, 1961). The choroidal melanocyte is congested with melanosomes, predominantly at the terminal Stage IV state of maturation (Fig. 4.2). Relatively minimal amounts of other cellular organelles, consisting of endoplasmic reticulum, Golgi apparatus, mitochondria, etc., exist in the cytoplasm of the choroidal melanocyte. In contrast, the RPE melanocytes are highly polarized cells approximately 14 mm in height and width. They form a single layer of cuboidal cells at the uveal/retinal junction and are connected to each other on their juxtaposed lateral surfaces by both elaborate terminal bar complexes, consisting of both

S Sinus

*

C *

RPE

R

A

B

P

Fig. 4.1. The back of the eye (A) consists of four major layers: the innermost neural retina (R), the retinal pigmented epithelium (RPE), the choroid (C), and the outermost sclera (S), as shown in the light micrograph cross-section of an avian eye. Melanocytes are conﬁned to the RPE and the choroid (B). The melanocytes of the choroid are heavily pigmented and multidendritic. They reside in the connective tissue component between the extensive sinus (sinus) and blood vessels (*) of the choroid. Melanocytes comprising the retinal epithelium have a relatively amelanotic cell body, with prominent nuclei (arrowheads) adjacent to the choroid, and numerous slender, pigmented dendrites (arrows) extending toward and interdigitating with the photoreceptor cells (P) (see also Plate 4.1A and B, pp. 494–495). Bar in A, 50 mm; bar in B, 15 mm.

93

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E

bv F

*

* N

M *

bv

RPE

zonula occludens and zonula adherens (Hudspeth and Yee, 1973), and desmosomes (Miki et al., 1975) (Fig. 4.3). Morphologically, these cells have a basal surface facing the uveal tract with numerous regular invaginations of the plasma membrane, called basal infoldings, extending 1 mm in length. Between these basal infoldings and the choroid of the uveal tract there exists: (1) an extremely thickened basement membrane (of approximately 2 mm) adjacent to the RPE melanocytes called Bruch’s membrane; and (2) an extensive anastomosing network of fenestrated blood vessels called the choriocapillaris adjacent to the stroma of the choroid (Fig. 4.3). Both these structures facilitate the transport of nutrients, waste, and presumably cytokines to and from the retina proper (Bill, 1975; Torczynski, 1982). Facing the photoreceptors of the retina, the RPE melanocyte apical surface has extensive slender cellular processes, the apical villi, that interdigitate between the photoreceptor cells (Fig. 4.3). These villi impart two basic functional roles unique to the RPE 94

Fig. 4.2. The choroid is the connective tissue spanning the sclera and the retinal pigmented epithelium (RPE). It contains numerous ﬁbroblasts (F) and dendritic melanocytes (M) that have cytoplasm congested with Stage IV melanosomes. Also distributed throughout the choroid is extracellular ground substance (*), nerve bundles (N), collagen (arrows), elastin, and endothelial cells (E) lining the extensive blood vascular network (bv) of the choroid. Bar = 3.0 mm.

melanocytes. First, the apical processes can lengthen or contract in correlation with the increased or decreased amount of light that enters the orbit, although it has been suggested that the melanosomes within the villi, and not the villi themselves, are translocated in response to light (Cohen, 1992). It is presumed that this translocation functions to coordinate the surface area of the photoreceptor cells available to receive photon activation under differential lighting conditions in the environment. The mechanism regulating the movement of the apical process of the RPE melanocyte or the melanosomes within the villi is unknown but is thought to be mediated through actin ﬁlaments (Burnside and Laties, 1976). Second, the RPE melanocytes also somehow facilitate the regeneration of rhodopsin, the photochemical responsible for transducing the visual signals (Dowling, 1960; Hubbard and Colman, 1959). In addition, the apical processes also participate in the phagocytosis of the eventually shed tips of the photoreceptor cells (Hogan et al., 1974) for the subsequent degradation of

EXTRACUTANEOUS MELANOCYTES

cap

*

* G RPE

PC

Fig. 4.3. The retinal pigment epithelium (RPE) is a single layer of polarized cuboidal melanocytes positioned between the choroid and the photoreceptor cells (PC) of the neural retina. Elaborate specializations exist at the RPE/choroid junction, which consist of the choriocapillaris (cap) and Bruch’s membrane (brackets) of the choroid and the basal infoldings (asterisks) of the RPE. At the RPE/neural retina junction, the RPE melanocytes exhibit numerous slender apical processes containing melanosomes (arrows), which interdigitate between the rod and cone photoreceptor cells (PC). The cell body of the RPE melanocyte is also relatively polarized, in that Stage IV melanosomes are predominantly restricted to the apical half of the cell. Also distributed throughout the cytoplasm of the RPE melanocyte are the Golgi apparatus (G), lipofuscin granules (arrowheads), and mitochondria (open arrows). Bar = 1.5 mm.

this material in the cytosol of the RPE melanocyte. The RPE melanocytes thus contain an elaborate lysosomal system for this purpose (Feeney, 1973). Abundant in the RPE melanocyte are residual bodies, the end-stage of the lysosome-based process of phagosomal degradation, which have been called lipofuscin granules (Weiter et al., 1986; Wing et al., 1978). These granules can be distinguished from the melanosomes of the RPE melanocytes in that the former are very irregular in shape and their inclusions predominantly consist of laminated whorls representing degraded lipids (Feeney-Burns, 1980). However, melanosomes can occasionally be incorporated into these residual bodies. The total amount of melanin eventually produced by the choroidal melanocytes and the RPE varies markedly among individuals. However, it has been shown that between races

this variation is not signiﬁcantly different for RPE melanin. In contrast, choroidal melanin is approximately twice as dense in African Americans as opposed to Caucasians (Geeraets et al., 1962; Hunold and Malessa, 1974; Weiter et al., 1986). The morphology of the melanosomes synthesized by the melanocytes of the uvea and the RPE can also be distinguished from each other. The shape of melanosomes in the uveal melanocyte is oval to spherical and generally 0.3 to 0.4 mm in diameter (Feeney and Hogan, 1961) but can be as large as approximately 1 mm in diameter (Boissy et al., 1988). In contrast, the melanosomes in the RPE melanocyte are extremely elongated and rod shaped ranging from 1.0 to 2.5 mm in length and 0.5 to 1.5 mm in width (Feeney-Burns, 1980; Kaczurowski, 1962). These distinctive RPE melanosomes are primarily localized toward and within the apical villi. However, spherical melanosomes ranging dramatically in diameter from 0.5 to 4.5 mm can also exist in the more perinuclear areas of the RPE (Kaczurowski, 1962). Another characteristic unique to the ocular melanocyte, in contrast to follicular and interfollicular cutaneous melanocytes, is their permanent halt in melanin synthesis and retention of melanosomes that occur shortly after birth of the organism. Melanization in the melanocytes of both the uveal tract and the RPE is transient, occurring only for a short period of time after initial differentiation in the target site (Boissy et al., 1988; Feeney-Burns and Mixon, 1979; Hu and Montagna, 1971). This contrasts markedly with melanocytes of the interfollicular epidermis which continuously synthesize melanin and transfer melanosomes to neighboring keratinocytes throughout the life of the organism (see Chapters 8 and 9). Hair follicle melanocytes also continually synthesize melanin and transfer melanosomes during their differentiated state, which occurs intermittently during the anagen cycles of hair growth (see Chapter 3). The transcription of melanocyte-speciﬁc genes responsible for the synthesis of melanin, and the subsequent production of melanin, has been documented in ocular melanoblasts during embryogenesis. Histologically, melanin synthesis (Mund et al., 1972; O’Rahilly, 1975) appears ﬁrst in the RPE melanocytes at 10–12 weeks of gestation in the human embryo and, as demonstrated in primates, subsequently in the uveal melanocytes at about 28–36 weeks in equivalent human gestation (Endo and Hu, 1973; Hu et al., 1973). In the mouse, transcripts for Dct/Tyrp2 were initially identiﬁed in presumed RPE melanocytes at the optic cup at 9.5 days post coitum (dpc) and in presumed uveal melanoblasts occurring in the mesenchyme immediately outside the developing optic vesicle at 11.5 dpc (Steel et al., 1992). By birth, both groups of ocular melanocytes contain numerous Stage IV melanosomes congestively packed within their cytoplasm. However, melanin synthesis is still occurring at this time. The increase in melanin (Matheny and Dolan, 1975; Pierro, 1963), expression of tyrosinase activity (Miyamoto and Fitzpatrick, 1957), DOPApositive reactivity in the trans-Golgi network (Endo and Hu, 1973), and the presence of tyrosinase, Tyrp1, and Dct/Tyrp2 (Chiu et al., 1993) have been demonstrated in neonatal eyes 95

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of various species. These criteria of active melanization disappear permanently within days or weeks of birth leaving both groups of ocular melanocytes melanogenically inert for the rest of the life of the individual (Hearing et al., 1973; John and Fitzpatrick, 1972). However, it has been reported that eye color can darken with age, suggesting that melanogenesis may be reactivated in the melanogenically silent ocular melanocytes (Carino et al., 1994; see also review by Schraermeyer, 1993). It has recently been demonstrated that cultured human ocular melanocytes can be induced by alpha melanocyte-stimulating hormone (a-MSH) under certain conditions to increase DOPA oxidase activity (Smith-Thomas et al., 2001). Certain pathologic and therapeutic conditions also demonstrate that ocular pigmentation can be regulated postnatally. Many human ocular diseases have been linked to ocular pigmentation or directly attributed to the abnormalities of ocular melanogenesis, as exempliﬁed by uveal melanoma, macular degeneration, senile cataract, Fuch’s heterochromic uveitis, and pigmentary glaucoma (Aggarwal et al., 1994; Cruickshanks et al., 1993; Regenbogen and Naveh-Floman, 1987; Scotto et al., 1976; Spector, 1984; Weiter et al., 1985). There is also evidence implicating a physiological association between innervation and pigmentation in the eye. Autonomic nerves innervate the eye in general (Laties, 1974; Laties and Lerner, 1975; Palumbo, 1976). More speciﬁcally, both adrenergic and cholinergic nerves frequently juxtaposition against melanocytes of the iridial stroma (Calhoun, 1919; Laties, 1974; Laties and Lerner, 1975; Ringvold, 1975). This iridial innervation appears to have inﬂuences on the regulation of ocular pigmentation, especially iris color. Cervical sympathectomy in various animal models has resulted in a marked decrease in iris coloration (Angelucci, 1893; Dryja and Albert, 1980; Giles and Henderson, 1958; Krebs et al., 1989; Laties, 1974). In humans, Horner’s syndrome results from a congenital or acquired unilateral paralysis of the cervical sympathetic nerves. This syndrome can frequently present with hypopigmentation of the iris associated with the lesional side (Calhoun, 1919; Weinstein et al., 1980). A postmortem electron microscopic analysis of the irises from a patient with Horner’s syndrome demonstrated that, in conjunction with a depletion of sympathetic nerve ﬁbers, stromal melanocytes were reduced in numbers with a few morphologically normal melanocytes remaining (McCartney et al., 1992). This suggests that iris stromal melanocytes depend on the sympathetic nerve supply to survive. Furthermore, epidermal melanocytes were also demonstrated electron microscopically to be innervated by neurons (Hara et al., 1996). Cultured uveal melanocytes demonstrate stimulated growth and melanogenesis upon exposure to epinephrine, isoproterenol, salbutamol, and metaproterenol, adrenergic agonists that can activate beta(2)-adrenoceptors (Hu et al., 2000). Prostaglandins have been a successful therapeutic treatment for intraocular pressure in glaucoma (Bito et al., 1987, 1993). Latanoprost, a phenyl-substituted analog of prostaglandin F2a administered as eye drops, has been demonstrated to darken the eye color of monkeys (Selen et al., 1997) and glaucoma 96

patients (Wistrand et al., 1997). Recently, it has been demonstrated that latanoprost can induce tyrosinase gene transcription in cultured iridial melanocytes (Lindsey et al., 2001; Stjernschantz et al., 2000).

Variation in Morphology/Function of Uveal and RPE Melanocytes Uveal Tract The uveal tract consists of the choroid, the ciliary bodies, and the iris. It is predominantly composed of the choroidal component that encases the entire posterior orbit of the eye. As the choroidal layer approaches the anterior, frontal area of the orbit, it transforms ﬁrst into the stromal component of the ciliary bodies and then into that of the iris, the anterior termination of the uveal tract. In a similar fashion, the RPE melanocyte layer is primarily found in the posterior orbit juxtaposed with the choroid. It also consists of anterior localized components that become the covering layer of the ciliary bodies and the inner or posterior surface of the iris. The bi- and/or multidendritic melanocytes of the choroid exhibit changes in their morphology as they reside in the stromal part of the ciliary bodies and the iris. At the base of the ciliary bodies, which is directly contiguous with the anteriormost area of the choroid, the melanocytes become relatively adendritic, in that they appear as large spherical cells congested with melanosomes. These spherical uveal melanocytes of the ciliary body are densely packed (Fig. 4.4). The ciliary bodies are primarily composed of processes that

C

NPE

PE CB

IRIS

Fig. 4.4. The pigmented tissues of the posterior chamber of the eye, i.e. the choroid and retinal pigmented epithelium (asterisk), are modiﬁed apically, ﬁrst into the ciliary bodies (CB) and then into the iris (I). The ciliary body is a pleated structure with a heavily pigmented uveal stroma (arrow) lined by an epithelial bilayer of proximal pigment cells (PE) and distal nonpigmented cells (NPE). The iris lies beneath the cornea (C) and is composed of an anterior (outer) face containing a heavily pigmented stroma continuous with the uvea of the ciliary bodies, and a posterior (inner) face containing a bilayer of pigmented melanocytes continuous with the RPE layer of the ciliary bodies. Bar = 100 mm.

EXTRACUTANEOUS MELANOCYTES

extend into the vitreous body forming an extensively pleated, cylindrical ring-like structure anchoring the zonular apparatus (i.e. system of 10-nm ﬁbers) responsible for accommodation of the lens during focusing (Coleman, 1970). The melanocytes distributed throughout the stroma of this part of the ciliary body become dendritic and resemble the melanocytes of the stroma of the choroid. The morphology of the melanosomes is identical to those of the choroidal melanocyte. The uveal tract leaves the ciliary bodies and continues on as the anterior stroma of the iris (Fig. 4.4) forming a circular ring of tissue (for review, see Miller, 1985). The iris, which is approximately 12 mm in thickness, marks the most frontal region of the posterior chamber of the eye and forms a diaphragm or wall separating it from the anterior chamber of the orbit. The most anterior surface of the iris is covered by a discontinuous layer of ﬁbroblasts. Immediately beneath this layer of ﬁbroblasts is a population of stromal melanocytes with dendrites orientated parallel to the iris surface. The remainder of the uveal component of the iris, called the iris stroma, contains numerous melanocytes with dendrites that are more extensively branched and more irregularly oriented. The melanosomes occurring in the melanocytes of the iris stroma, like those throughout the rest of the uveal tract, are oval to spherical in shape, measuring between 0.25 and 0.75 mm in diameter (Zinn et al., 1973). However, the size of these melanosomes varies depending on the genetic makeup of the color of the iris (see below). Like the choroid and ciliary body stroma, the iris stroma also contains an extensive amount of ﬁbroblasts, blood vessels, nerves, and collagen ﬁbers.

Clump Cells of Koganei Unique to the iris stroma is a cell type called the clump cell of Koganei (Wobmann and Fine, 1972). This spherical cell, much larger than the melanocyte, has a cytoplasm congested with many membrane-bound vesicles varying dramatically in size and containing primarily melanosomes with some intermittent granular and/or particular material. These giant pigmented cells have been proposed to be resident macrophages that have phagocytized a large number of melanin granules (Wobmann and Fine, 1972). Macrophages that appear to be precursors of clump cells of Koganei can occasionally occur on the anterior surface of the iris. These cells increase both in number and in content of engulfed melanosomes either during aging or in some pathologic conditions (Boissy et al., 1987; Hogan and Zimmerman, 1962; Wobmann and Fine, 1972). The most posterior face of the iris is lined with a double layer of melanocytes derived from the anteriormost continuation of the RPE layer as detailed below.

Retinal Pigment Epithelium Similar to the uveal tract, the RPE melanocyte also exhibits morphologic changes as it progresses anteriorly along and in juxtaposition with the uveal layer. Covering the ciliary bodies, the RPE becomes a bilayer called the ciliary epithelium (i.e.

ciliary RPE) lined on both surfaces by a basement membrane (Fine and Zimmermann, 1963; Straatsma et al., 1968). The cells of the outermost layer are relatively devoid of melanosomes and are called the nonpigmented epithelial cells (NPE) (Fig. 4.4). In contrast, the innermost layer contains numerous melanosomes and is called the pigmented epithelium (PE) (Fig. 4.4). These two cell layers of the ciliary epithelium are interjoined by numerous junctions including gap junctions, desmosomes, and tight junctions (Raviola and Raviola, 1978). The melanosomes of the PE resemble those of the RPE except that a higher percentage of them are more spherical and less elliptical. However, as a group, they are much larger than the melanosomes in the melanocyte of their stromal counterpart (Streeten, 1982). The RPE layer extending anteriorly toward the iris eventually forms the posterior leaf of this tissue and is speciﬁcally called the iris pigment epithelium (IPE) (for review, see Miller, 1985). The IPE, like that of the ciliary epithelium, is a double layer of cuboidal cells rather than a single layer. The posterior layer of the IPE is composed of heavily pigmented cuboidal melanocytes with a typical basement membrane on the posterior surface separating the cells from the vitreous chamber of the eye. The lateral surfaces of the posterior IRP cells are extensively connected to each other by lateral infoldings, desmosomes, and tight junctions. The posterior or basal surface manifests areas of infoldings, whereas the anterior or apical surface expresses numerous desmosomes and tight junctions adhering these cells to the anterior layer of the IPE. The anterior layer of the IPE is composed of an extremely unique type of melanocyte that expresses numerous myoﬁlaments and is called the anterior epithelium–dilator muscle layer. The elongated melanocytes exhibit striking polarity and measure approximately 12.5 mm in height. The basal cytoplasm contains the nucleus and most of the heavily pigmented melanosomes. In contrast, the apical portions, which are wavy or spindle-shaped and interwoven with the apical portions of neighboring melanocytes, are relatively devoid of melanosomes and contain the classical arrangement of myoﬁlaments. When these myoﬁlaments induce contraction of the apical processes of these IPE cells, the iris layer is retracted and the pupil dilates (Salzmann, 1912; van Alphen, 1963, 1979). The contractile properties of the apical processes of the IPE cells resemble those of the apical processes of the RPE cells interdigitating with the photoreceptor cells of the retina as described above. Regulation of this contractile property appears to be under direct sympathetic stimulation by either neighboring neurons or humoral adrenergic substances such as norepinephrine and adrenal catecholamines (Miller, 1985). Therefore, it has been proposed that the ocular melanocyte must express functional receptors responsive to various neurokines that inﬂuence this contractile property (Bonvallet and Zbrozyna, 1963; Miller, 1985; van Alphen, 1963, 1979) and possibly melanogenesis as well (Laties, 1974; McCartney et al., 1992), as described earlier in this chapter. Finally, the melanosomes in the melanocytes of both layers of the IPE are relatively elongated like those distinctive of the rest of the RPE 97

CHAPTER 4

and are on average 0.8 mm in diameter and 2.5 mm in length (Zinn et al., 1973).

Eye Color Mammalian The color of human irises exhibits a transitional range from light (blues) to medium (greens/hazels) to dark (browns) and was demonstrated at the turn of this century to be under genetic regulation (Davenport and Davenport, 1907). The ultimate inherited color of an iris appears to be under the inﬂuence of both the amount of pigmentation in and the size of melanosomes within the stroma of the iris, and not by the number, density, and/or shape of the melanocytes in the stroma nor the melanosomes or melanocyte components of the IPE (Imesch et al., 1996; Wilkerson et al., 1996). The number, size, and pigmentation of the melanosomes produced in both cuboidal layers of the IPE appear to be constant and maximal regardless of the iris color (Feeney et al., 1965; Fine and Yanoff, 1979). In contrast, initial investigations demonstrated that variation in pigmentation of the anterior stroma layer correlated with eye color (Fuchs, 1913; Wolfrum, 1922). It was originally proposed that the number of melanocytes in the iris stroma increased as the color of the iris darkened (Dietrich, 1972). However, more recent quantitative analyses have demonstrated that the number of stromal melanocytes does not differ with eye color (Eagle, 1988; Wilkerson et al., 1996). Instead, quantitative differences in the density (Eagle, 1988; Imesch et al., 1996) and size (Eagle, 1988) of melanosomes do vary, in that stromal melanocytes of darker colored eyes contain larger and more numerous melanosomes per cytoplasmic area than lighter colored eyes. However, it has not yet been determined whether the quantity or quality of melanin within the melanosomes also varies with iris color. It is possible that the amount of eumelanin vs. pheomelanin or the ratio of brown-soluble vs. black-insoluble forms of eumelanin varies between different colored irises. It has recently been demonstrated that the ratio of eumelanin to pheomelanin in ocular tissue does not differ in blue or brown eyes (Menon et al., 1992) or in bovines of different coat colors (Dryja et al., 1979). The question of variation in melanogenesis correlating with eye color is being readdressed with the indication that both amount and type of melanin varies with different iris coloration (Sarna and Wielgus, 2004). Therefore, it is the function of melanocytes once established in the iridial stroma, and not the embryonic development of the melanocyte population in the eye, that determines iris color. Ultimately, the increased amount of backscattered reﬂective light occurring as a consequence of less melanosomal density in blue eyes, as opposed to more light absorption and relatively less reﬂection of light due to the relative higher density of melanosomes in brown eyes, is principally responsible for the color of the iris one observes (Miller, 1978).

Nonmammalian While the color of the iris of most birds and mammals is brown to black, there are notable exceptions of yellow, red, 98

RP

M

M

Fig. 4.5. The iris of the ground dove contains iridophores, congested with spaces representative of reﬂecting platelets (RP), residing next to myocytes (M) of the stroma. Bar = 1.0 mm (micrograph courtesy of Joseph Bagnara).

or orange irides, especially among avian species. This is not so surprising as the integument of birds and lower vertebrates also exhibits yellow, red, and orange as a result of the involvement of other neural crest-derived chromatophores, i.e. iridophores/leukophores and xanthophores/iridophores, in addition to melanophores (see Chapter 2). A study of the irides of two species of ground doves with yellow/orange and deep red iris color (Ferris and Bagnara, 1972) revealed the presence of true iridophores and their constituent reﬂecting platelets (Fig. 4.5). It is presumed that these capillary-associated iridophores function to enhance the red color stemming from the capillaries during vasodilation (Chiasson et al., 1968). No evidence of yellow chromatophores with their constituent pterinosomes was seen in the irides of ground doves, although the possible presence of pteridines was suggested by the ﬂuorescence exhibited by iris extracts. A study of the great horned owl iris with yellow irises by Oliphant (1981) revealed the presence of both pterinosome-like organelles and pteridines of the type usually found in xanthophores and erythrophores of cold-blooded vertebrates. It was suggested that crystalline deposits of pteridines were present in the reﬂecting platelets

EXTRACUTANEOUS MELANOCYTES

of the iris, a fact later shown to be true for the reﬂecting platelets of frog iridophores (Bagnara et al., 1988). Observations on the irides of other avian species revealed the presence of both xanthophore and iridophore pigments (Oliphant, 1987a, b, 1988) and, later, Hudon (1994) extended the list of avian species possessing these pigments and demonstrated that irides of some birds possess unique pigments. One of these, pterorhodin, is a pteridine dimer that has been found in the chromatophores of only one vertebrate group, a subfamily of tree frogs (see Chapter 2). Thus far, mammalian irides have not been studied from the aspect of xanthophore or iridophore presence, although some regular and elongated organelles described from the cat iris (Tousimis, 1963) may represent the presence of iridophores.

Otic Melanocytes At the outset, we recommend the following excellent literature for more substantial information on the development and anatomy of the ear (Anson and Davies, 1980; Donaldson and Miller, 1980) and its melanocyte component (Cable and Steel, 1991; Conlee et al., 1989; Meyer zum Gottesberge, 1988a; Tachibana, 1999). Melanocytes have been identiﬁed in various sites of the ear. Primarily, they are localized throughout the stria vascularis and the modiolus of the cochlea, but they also exist in the vestibular organ composed of the utricle, the ampullae, the semicircular ducts, and the saccule as well as in the endolymphatic sac (Meyer zum Gottesberge, 1988a). In all cases, the melanocytes are extensively dendritic, heavily pigmented, and basically reside beneath the epithelial layer of each respective area. However, in the stria vascularis, the melanocytes become dramatically interdigitated with the extensive basolateral plasma membrane infoldings of the marginal (i.e. epithelial) cells (see below) (Cable and Steel, 1991). Much of the structural/functional analyses of otic melanocytes have almost exclusively focused on the melanocytes residing in the stria vascularis of the murine and/or chinchilla models. Therefore, much of the following discussion of otic melanocytes pertains to this subpopulation. Although the neural crest origin of otic melanocytes has not been as rigorously determined, using labeling and/or transplantation studies, as that of cutaneous and ocular melanocytes, it has nonetheless been proposed, primarily on account of association, that the otic melanocytes are of neural crest origin (Hilding and Ginzberg, 1977). The most convincing data implicating the neural crest as the source from which otic melanoblasts arose come from observations of mice mutant at the W locus. The W gene encodes the c-kit tyrosine kinase receptor that regulates the embryonic development of various cell types, speciﬁcally neural crest-derived melanocytes of the skin and uveal tract (Geissler et al., 1988; Hilding and Ginzberg, 1977; Mackenzie et al., 1997). Concomitantly, mutant W mice have a loss of melanocytes in the ear (Deol, 1970a; Schrott and Spoendlin, 1987; Steel et al., 1987). Studies on mice mutant for various alleles of the W locus demonstrated a halt in the expression of Dct/Tyrp2-positive

cells (i.e. presumptive melanoblasts) at a migratory site outside the developing ear (Cable et al., 1995), similar to the defective survival and migration of neural crest-derived melanoblasts observed in KitW-v/KitW-v mice during embryonic development using a different melanoblast marker (Mackenzie et al., 1997). The similarity between the in vivo effects of Kit mutations upon melanoblasts migrating to both the integument and the otic vesicle, together with the lack of an apparent effect of Kit mutations upon RPE melanocyte development, provides important correlative evidence for the neural crest origin of otic melanocytes. Interestingly, Mitf, but not Sox10, is expressed in strial melanocytes in immediately postnatal mice, despite the fact that Sox10 regulates Mitf expression during melanocyte development (Bondurand et al., 2000; Potterf et al., 2000), suggesting that this regulatory event in otic melanocytes may be speciﬁc to earlier developmental stages of those cells (Watanabe et al., 2002). Minimal amounts of melanin synthesis have occurred in the stria vascularis by birth (Fig. 4.6). During the subsequent weeks, melanogenesis dramatically increases until the otic melanocytes appear congested with melanosomes (Cable and Steel, 1991), similar to the process occurring in uveal and dermal melanocytes. Both eumelanin and pheomelanin appear to be synthesized by these otic melanocytes (Inoue et al., 1984). However, DOPA-reactive products in the trans-Golgi network, indicating active synthesis of tyrosinase, continue to be detectable well into adult life (Cable and Steel, 1991; Escobar et al., 1995), contrasting with the halt in tyrosinase synthesis postnatally that occurs in uveal and dermal melanocyte populations (Chiu et al., 1993). Although there is no concomitant synthesis of premelanosomes at this time, the target site and ultimate function of the tyrosinase-like enzyme in otic melanocytes of the adult remain unknown. Interestingly, it has also been demonstrated that the amount of otic melanosomes can increase in adult animals after exposure to impulse noise (Gratton and Wright, 1992) and experimentally induced endolymphatic hydrops (Meyer zum Gottesberge, 1986). The stria vascularis of the ear continues to develop after birth with extensive restructuring of the marginal, intermediate (i.e. melanocytes), and basal cell layers. As a result, there is a coordinate development of the endocochlear potential of the ear and the interdigitating of the melanocytes and the elaborate basal processes of the marginal cells (Cable and Steel, 1991). At this time, the melanocytes in the intermediate cell layer are called light cells. Morphologically, they have an extensive amount of organelles involved in protein synthesis (i.e. RER and Golgi apparatus) and numerous melanosomes individually distributed throughout the cytosol. Eventually, dark intermediate cells, which exhibit relatively no cytoplasmic organelles except for complexed, acid phosphatasepositive melanosomal complexes plus a pyknotic nucleus, appear in the same vicinity. These dark cells are presumed to be a terminal stage toward which the light intermediate melanocytes eventually progress (Cable and Steel, 1991). This is very unique, as no other subpopulation of melanocytes 99

CHAPTER 4

* *

M

E IV III

II E

throughout the body ever naturally progresses to such a morphologically necrotic-like state. Many functions for otic melanocytes in the physiology of the ear and hearing have been proposed and debated. These putative roles include: (1) the embryonic development of the ear and hearing; (2) the maintenance of endocochlear potential; and (3) the prevention of noise- and/or toxin-induced hearing loss. In general, if otic-destined, neural crest-derived melanoblasts do not migrate successfully into the ear or fail to survive in that location as a function of mutations in spotting genes such as Kit (Schrott and Spoendlin, 1987; Steel et al., 1987), KitL (Mayer and Green, 1968), or Mitf (Deol, 1970b, 1971; Green, 1989), hearing loss occurs. Similarly, deafness in humans that occurs in the majority of patients with Waardenburg syndrome (Hageman and Delleman, 1977; Read and Newton, 1997) and occasional variants of piebaldism (Giebel and Spritz, 1991; Hulten et al., 1987) has also been associated with the lack of normal otic melanocyte development. In KitW-v/KitW-v murine mutants, absence of strial melanocytes results in reduced interdigitations between marginal cells and basal cells, changes that can be observed as early as 6 days after birth (Steel and Barkway, 1989). More extensive cellular and structural degeneration of the cochlea, apparent after 6 weeks of age, occurs in other coat color mutants that lack cochlear melanocytes (Deol, 1970a). In addition, the morphology of the marginal cells is also compromised in cases where melanocytes are successfully established in the inner ear but remain amelanotic, i.e. in albinism (Conlee et al., 1986; Conlee and Bennett, 1993). Albinism also presents with abnormal brainstem auditory evoked responses associated with a reduction in neuronal size of the superior olivary complex (Baker and Guillery, 1989; Conlee et al., 1984, 1986; Creel et al., 1980, 1983). This is reminiscent of the neural alterations in the visual system that 100

Fig. 4.6. Otic melanocyte in the apical coil of the cochlea in an 18.5-day mouse fetus. The cell body of the melanocyte (M) resides among the epithelial cells (E) of the developing stria vascularis. Developmental stages (II, III, and IV) of melanosome biogenesis are present in the melanocyte. A cytoplasmic process of the melanocyte (arrowhead) appears to extend into the underlying mesenchyme (asterisk) through gaps (arrows) in the basal lamina (open arrows), perhaps indicating melanocyte migration from the mesenchyme into the epithelium. Continuity of this process with the cell body of the melanocyte was conﬁrmed in serial sections (inset). Bar = 0.5 mm (micrographs courtesy of Rick Friedman and Emma Lou Cardell).

also correlate with the absence of melanin in albino models (see previous section in this chapter). In summation, the presence of pigmented melanocytes appears to be essential for the complete development of the marginal cells and the otic centers of the brain. The endocochlear potential, a measurement that reﬂects the ionic and electrochemical gradients within the ﬂuids of the inner ear responsible for transducing sound waves to the hair cell receptors (Bosher et al., 1973), is also affected in aberrant pigment systems. Evidence supporting the importance of melanocytes to the generation of the endocochlear potential has also resulted from the analysis of murine coat color mutants. In most KitW-v/KitW-v mice, the development of an endocochlear potential, which normally occurs gradually over the course of the ﬁrst two neonatal weeks, does not occur at all in the large majority of homozygous mutants that contain no strial melanocytes (Steel and Barkway, 1989). In addition, the extent to which the endocochlear potential is suppressed in development correlates with the extent of otic depigmentation associated with the various W alleles (Cable et al., 1994). The endocochlear potential in albinos is also reported to be altered (Conlee and Bennett, 1993; Conlee et al., 1991), although this has been contested (Bock and Steel, 1984). Therefore, hearing loss in these mutant pigmentary systems results directly from the absence of an endolymph mediator system. A combination of electrophysiologic, immunolocalization, and knockout mouse studies has demonstrated that the melanocyte, or intermediate cell, in the cochlea expresses the potassium ion channel directly responsible for EP generation. The depression of the EP by Ba2+ suggested that an inwardly rectifying K+ (Kir) channel located in the stria vascularis was responsible for the EP. A screen conducted of the stria vascularis for expression of various Kir channels revealed that only

EXTRACUTANEOUS MELANOCYTES

one, Kir4.1, was expressed in this tissue (Hibino et al., 1997). Although the cell type expressing Kir4.1 was originally in question, whole-cell patch clamp experiments described Ba2+responsive, inwardly rectifying K+ currents across the cell membranes of dissociated strial intermediate cells (Takeuchi and Ando, 1998), leaving open the possibility that the intermediate cell is responsible for generating the EP because of Kir4.1 expression. A subsequent study using an anti-Kir4.1 antibody in conjunction with immunoﬂuorescence of dissociated strial cells and immunogold electron microscopy localized Kir4.1 to the intermediate cell membrane (Ando and Takeuchi, 1999). Finally, the lack of EP observed in mice lacking the Kir4.1 channel due to a homozygous knockout mutation conﬁrmed the important role of this channel in supplying the electromotive force required for EP generation (Ando and Takeuchi, 1999). Melanin in otic tissues has been proposed also to protect against the loss of hearing resulting from noise- and/or toxininduced trauma. Susceptibility to noise-induced hearing loss has been demonstrated in some animals and humans with albinism (Barrenas and Lindgren, 1990; Conlee et al., 1986, 1988; Hood et al., 1976). However, the baseline hearing capabilities in albinos do not appear to be compromised (Harrison et al., 1984; Versnel et al., 1990; Wasterstrom, 1984). The presence of otic melanin has correlated with susceptibility (Comis and Leng, 1980; Wasterstrom, 1984), as well as resistance (Conlee et al., 1991), to ototoxicity. Therefore, the role of melanin in the ear is complex. It can absorb excess sound waves and toxins to prevent the subsequent damage to hair cells. On the other hand, melanin can promote ototoxicity depending on the circumstances (Dencker et al., 1973; Lindquist, 1973; Lindquist and Ullberg, 1972; McGinness et al., 1974; Meyer zum Gottesberge, 1988b). Currently, it has been questioned whether the maintenance of melanocytes and pigment in the ear of adults is essential. Loss of melanocytes, which occurs in brown mice homozygous for the Blt allele at the gene encoding Tyrp1 (Cable et al., 1993) and human patients with vitiligo (Dereymaeker et al., 1989; Orecchia et al., 1989; Tosti et al., 1987), has been associated with the development of hearing loss. It has also been suggested that pigment loss in the ear in conjunction with aging has a cause and effect relationship with the eventual deterioration of otic activity (Dum, 1982; Rosen et al., 1962).

Melanocytes of Internal Body Sites Melanocytes in the Leptomeninges A focal area of the human brain, speciﬁcally the meninges covering the ventrolateral surfaces of the medulla oblongata and the upper cervical cord, appears to be macroscopically pigmented (Baader, 1935; Symmers, 1905). This pigmentation is the consequence of melanocytes within this tissue, as conﬁrmed by ultrastructural analysis (Goldgeier et al., 1984). Melanocytes residing in the meninges of the mouse have also been reported (Barden, 1986). In the adult human, melanocytes of the leptomeninges are multidendritic and congested with predominantly Stage IV melanosomes. The

presence in the cytoplasm of relatively few early-stage melanosomes and premelanosomes, as well as the absence of histochemically induced DOPA-positive reaction product in the trans-Golgi network, suggest that in the adult these cells are melanogenically inactive (Goldgeier et al., 1984), similar to the melanocytes of the dermis, the uveal tract, and the RPE. It should be noted that the pigment (i.e. melanin) of the melanocytes in the leptomeninges differs from the dark pigment found within neurons throughout the brain. The latter is classiﬁed as neuromelanin and is thought to be autooxidative product(s) of dopamine generated by the neuronal synthesized tyrosine hydroxylase molecule (see Breathnach, 1988; Enochs et al., 1993, 1994; Odh et al., 1994; Zecca et al., 1992). The function of the melanocytes conﬁned to the leptomeninges is not clear. Of interest, it has been speculated that these melanocytes may be affected in patients with severe vitiligo (i.e. the Vogt–Koyanagi–Harada syndrome) and that their loss may correlate with the aseptic meningitis component of this syndrome (Goldgeier et al., 1984; Nordlund et al., 1980). Additionally, aberrant proliferation of these melanocytes during embryonic development, usually associated with the presence of a large congenital melanocytic nevus, may result in the condition neurocutaneous melanosis (Marghoob et al., 2004).

Melanocytes in the Harderian Gland The harderian gland is a secretory structure located at the medial back of the optic orbit with a duct that connects it to the nictitating membrane of the eyelid (Payne, 1994). It exists in all groups of terrestrial vertebrates but is absent in some types of mammals (i.e. bats, cows, and horses), terrestrial carnivores, and higher primates including man. Structurally, the gland is composed of tubules of simple columnar epithelial cells surrounded by myoepithelial cells, and it primarily synthesizes lipids, porphyrins, and indoles. Many putative and diverse functions have been hypothesized for this secretory gland (Payne, 1994). Melanocytes have been identiﬁed in the harderian glands of mice (Shirama et al., 1988), gerbils (Johnston et al., 1983), and frogs (Di Matteo et al., 1989). These melanocytes are primarily localized throughout the connective tissue capsule surrounding the harderian gland and intermittently among the connective tissue septa invaginations into the gland that separates the stroma into lobules (Fig. 4.7). Like the melanocytes of the leptomeninges, the melanocytes in the harderian gland of an adult animal are multidendritic, melanogenically silent, and congested with fully pigmented melanosomes (Fig. 4.7). Harderian glands ultrastructurally observed from the murine hypopigmented mutants ruby eye (Ru/Ru) and dilute (di/di) exhibited relatively fewer melanocytes with a dramatic reduction in melanosomes distributed throughout the cytoplasm (R. E. Boissy and K. Moore, personal observations). The function of melanocytes in the harderian gland is unknown, and no dysfunction in any of the putative roles of the gland has been reported in association with the various murine hypopigmentary mutants. 101

CHAPTER 4

E

E A

with the connective tissue coverings of cartilage (perichondrium) and bone (periostium). In addition, melanocytes were prevalent in the gonads, the lungs, the thymus gland, the thyroid gland, the spinal ganglia, and the gray matter of the spinal cord, but absent from the liver, the pancreas, and the spleen (Hallet and Ferrand, 1984; Makita and Mochizuki, 1984; Makita and Tsuzuki, 1986; Stolle, 1968). This mesenchymal distribution of melanocytes is primarily the result of an autosomal dominant gene (P) that can be intensiﬁed by a sex-linked recessive gene (d) (Stolle, 1968). Current analysis of the migration pathway of neural crest-derived melanoblasts in Silkie embryos demonstrated that these precursors are not restricted to the dorsolateral pathway as in normal development, but also migrate aberrantly through the ventral pathway (Reedy and Erickson, 1995). This genetically controlled diversion of melanoblast migration appears to be under cues extrinsic to this neural crest derivative (Hallet and Ferrand, 1984; see also Chapter 5).

Perspectives

E

B Fig. 4.7. The harderian gland is composed primarily of epithelial cells (E) housing numerous and varied types of secretory granules. Melanocytes congested with Stage IV melanosomes exist occasionally within the connective tissue septa separating the lobes of the harderian gland (arrow in A) and prominently within the connective tissue capsule encasing the harderian gland (arrow in B). Bar = 5.0 mm.

Extracutaneous melanocytes found in the eye, ear, and various internal body structures are unique as a whole from epidermal melanocytes in that: (1) they synthesize melanosomes and melanin only during a short interval after arriving in their target tissue; and (2) they retain their melanosomes throughout the life of the organism as opposed to transferring them to neighboring cells. The general morphology and ultrastructure of these extracutaneous melanocytes and the speciﬁc variations that exist among them have been well documented in the literature. However, the role of these morphologically dormant melanocytes in their respective tissues has primarily been ascertained anecdotally to date. Fortunately, experimental analysis investigating the function of extracutaneous melanocytes is beginning to make a reappearance in the scientiﬁc literature. The advent of molecular genetic technology, and its combination with cellular and physiological approaches, will help to resolve the elusive function of these relatively ignored populations of melanocytes that have been retained through evolution.

Melanocytes in Mesenchymal Tissues There have been a few reports of melanocytes distributed throughout internal body sites, most notably related to the PET mouse (Reams, 1963) and the Silkie chicken (Stolle, 1968). In the PET mouse, melanocytes appearing multidendritic, heavily pigmented, and inactive are distributed throughout the connective tissues of multiple sites in the body cavity, including cartilage, bone, serosae, lungs, heart, gonads, etc. (Nichols and Reams, 1960; Rovee and Reams, 1964). The speciﬁc genetics of this undeﬁned strain of mouse was not clearly described; however, the inheritance of the mesenchymal pigmentation in the PET mouse appears to have been autosomal recessive in nature (Mayer and Reams, 1962; Reams, 1967). In the Silkie chicken, melanocytes were identiﬁed in the loose connective tissue throughout the body and associated 102

Acknowledgments We wish to thank Joseph Bagnara for providing information and illustrations in the section on nonmammalian eye color and Emma Lou Cardell for providing information and the illustration on otic melanocytes.

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Tosti, A., F. Bardazzi, G. Tosti, and L. Monti. Audiologic abnormalities in cases of vitiligo. J. Am. Acad. Dermatol. 17:230–233, 1987. Tousimis, A. J. Pigment cells of the mammalian iris. Ann. N. Y. Acad. Sci. 100:447–466, 1963. Tropepe, V., B. L. Coles, B. J. Chiasson, D. J. Horsford, A. J. Elia, R. R. McInnes, and D. van der Kooy. Retinal stem cells in the adult mammalian eye. Science 287:2032–2036, 2000. Udono, T., K. Yasumoto, K. Takeda, S. Amae, K. Watanabe, H. Saito, N. Fuse, M. Tachibana, K. Takahashi, M. Tamai, and S. Shibahara. Structural organization of the human microphthalmia-associated transcription factor gene containing four alternative promoters. Biochim. Biophys. Acta 1491:205–219, 2000. van Alphen, G. W. The structural changes in meiosis and mydriasis of the monkey eye. Arch. Ophthalmol. 69:802–814, 1963. van Alphen, G. W. Scanning electron microscopy of the monkey eye in meiosis and mydriasis. Exp. Eye Res. 29:511–526, 1979. Versnel, H., V. F. Prijs, and R. Schoonhoven. Single-ﬁbre responses to clicks in relationship to the compound action potential in the guinea pig. Hearing Res. 46:147–160, 1990. Wasterstrom, S.-A. Accumulation of drugs on inner ear melanin: Therapeutic and ototoxic mechanisms. Scand. Audiol. Suppl. 23:1–40, 1984. Watanabe, K., S. Inai, K. Jinnouchi, S. Bada, A. Hess, O. Michel, and T. Yagi. Nuclear-factor kappa B (NF-kappa B)-inducible nitric oxide synthase (iNOS/NOS II) pathway damages the stria vascularis in cisplatin-treated mice. Anticancer Res. 22:4081–4085, 2002. Weinstein, J. M., T. J. Zweifel, and H. S. Thompson. Congenital Horner’s syndrome. Arch. Ophthalmol. 98:1074–1078, 1980. Weiter, J. J., F. C. Delori, G. L. Wing, and K. A. Fitch. Relationship of senile macular degeneration to ocular pigmentation. Am. J. Ophthalmol. 15:185–187, 1985. Weiter, J. J., F. C. Delori, G. L. Wing, and K. A. Fitch. Retinal pigment epithelial lipofuscin and melanin and choroidal melanin in human eyes. Invest. Ophthalmol. Vis. Sci. 27:145–152, 1986. Wilkerson, C. L., N. A. Syed, M. R. Fisher, N. L. Robinson, I. H. L. Wallow, and D. M. Albert. Melanocytes and iris color: light microscopic ﬁndings. Arch. Ophthalmol. 114:437–442, 1996. Wing, G. L., G. C. Blanchard, and J. J. Weiter. The topography and age relationship of lipofuscin concentration in the retinal pigment epithelium. Invest. Ophthalmol. Vis. Sci. 17:601–607, 1978. Wistrand, P. J., J. Stjernschantz, and K. Olsson. The incidence and time-course of latanoprost-induced iridial pigmentation as a function of eye color. Surv. Ophthalmol. 41 (Suppl. 2):S129–138, 1997. Wobmann, P. R., and B. S. Fine. The clump cells of Koganei: A light and electron microscopic study. Am. J. Ophthalmol. 73:90–101, 1972. Wolfrum, P. Uber den Bau der Irisvorderﬂache des menschliche Auges mit verleichend anatomischen Bemerkungen. Graefes Arch. Klin. Exp. Ophthalmol. 183:317–333, 1922. Zecca, L., C. Mecacci, R. Seraglia, and E. Parati. The chemical characterization of melanin contained in substantia nigra of human brain. Biochim. Biophys. Acta 1138:6–10, 1992. Zinn, K. M., S. Mockel-Pohl, V. Villanueva, and M. Furman. The ﬁne structure of iris melanosomes in man. Am. J. Ophthalmol. 76: 721–729, 1973.

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The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

5

Regulation of Melanoblast Migration and Differentiation David M. Parichy, Mark V. Reedy, and Carol A. Erickson*

Summary 1 The discovery of the relationship between the neural crest and pigment cells is brieﬂy reviewed. 2 In all animals studied thus far, neural crest cells are known to migrate along speciﬁed pathways and to differentiate into a variety of cell types. These phenomena are directed by a variety of determining factors. 3 With regard to understanding pigment cell migration and differentiation, ﬁsh have recently contributed in a signiﬁcant way primarily because of the large number of mutants that have been isolated in zebraﬁsh and other species. Moreover, the clear embryos are amenable to experimental manipulation and visualization. 4 Amphibians have provided extensive opportunities to examine the movement and differentiation of a variety of pigment cell types, and in this case a number of mutants have been studied in some detail. However, this system has not been exploited to its full potential. 5 Although many reports of pigment variants exist in reptiles, little work has been done on developmental aspects of reptilian chromatophores. 6 Our understanding of the cellular and biochemical controls of pigment cell development has been greatly enhanced by work done in avian systems. Recently the development of methods to up- and downregulate gene expression in the avian embryo has contributed importantly to the understanding of melanogenesis in this system. 7 The many coat color mutants of mice provide extensive opportunities for better understanding the motive forces in pigment cell development. 8 A comparative evolutionary approach to studye pigment cell development will continue to be essential for understanding how such discrete cells (pigment cells) from a well-deﬁned progenitor cell population (the neural crest) have differentiated and evolved.

Historical Background Embryologists have long been interested in the mechanisms that produce the correct arrangement of cells and tissues in *Kenneth A. Mason and Sally K. Frost-Mason were also authors of the previous version of this chapter.

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the developing embryo. Two of the processes that drive this event, called pattern formation, are morphogenesis and differentiation. Morphogenesis refers to the generation of biologic structure by the rearrangement of cells or tissues. In many cases this is achieved by the active migration of cells from one location in the embryo to another. Differentiation, on the other hand, is the process by which cells acquire morphologic and biochemical characteristics distinct from other cells. Although morphogenesis and differentiation usually are studied as independent developmental events, in fact these two processes are highly interconnected. The development of the neural crest exempliﬁes this complex interplay between morphogenesis and differentiation. Neural crest cells are a population of cells found in all vertebrate embryos. Excellent introductions to the older literature on neural crest development can be found in Hörstadius (1950) and Le Douarin and Kalcheim (1999). The early studies demonstrated three important aspects of neural crest development. First, neural crest cells detach from the neuroepithelium shortly before or soon after fusion of the neural folds. Second, neural crest cells migrate extensively throughout the developing embryo. Third, the neural crest is not a terminally differentiated population of cells, but instead can give rise to a wide range of cell and tissue types. These include neurons and glial cells of the peripheral nervous system, the secretory cells of the adrenal medulla, certain cardiac structures, enteric ganglia, and connective tissues of the craniofacial skeleton. Pigment cells are another derivative of the neural crest. There are several types of pigment cell, also called chromatophores, in vertebrate ectotherms (ﬁshes, amphibians, reptiles) including: black, melanin-containing melanophores; yellow, pteridine-containing xanthophores; and silvery, purine-containing iridophores. In contrast, endotherms (birds and mammals) have a single type of skin pigment cell, the melanin-containing melanocyte. The study of pigment cell development in vertebrates has a rich historical tradition, and many older reviews provide access to these classical studies (e.g., Bagnara and Hadley, 1973; Le Douarin, 1982). The purpose of this chapter is to present an overview of the current state of knowledge on the development of neural crestderived pigment cells in vertebrate embryos. We begin with a discussion of themes in neural crest and pigment cell development common to all vertebrates. Because this textbook is organized around the concept of taxonomy, we have similarly organized this chapter by taxon. Thus, we summarize what is

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known about pigment cell development speciﬁcally within the different classes of vertebrates: ﬁshes, amphibians, reptiles, birds and mammals. The reader is encouraged to note the similarities and differences in the control of pigment cell development across these major phylogenetic groups. Although mathematical modeling is one technique that has been employed to study the generation of pigment patterns in vertebrates (reviewed in Murray, 1990), we approach the topic from a molecular and cellular perspective, and thus do not discuss any of these models speciﬁcally. Instead, our discussion is framed around the subtopics of morphogenesis and differentiation. Morphogenesis and differentiation are treated separately because most empirical studies to date have focused principally on one process or the other. Nevertheless, there is clear evidence that these two events are closely linked during pigment cell development (reviewed by Erickson and Reedy, 1998). Additionally, we discuss the role that pigmentation mutants have played in elucidating the molecular control of pigment cell development. In some animal models, such as birds and amphibians, pigment mutants have been useful but not the primary means of understanding melanogenesis. In the case of ﬁshes and to a lesser extent mammals, most of what we know about pigment patterns has been derived from mutants and these are discussed extensively throughout the topics of morphogenesis and differentiation. Finally, we discuss major unresolved questions and suggest future research directions in the ﬁeld.

Current Concepts: Overview of Neural Crest and Pigment Cell Development An understanding of the mechanisms that control neural crest and pigment cell morphogenesis is predicated on knowledge of the precise pathways these cells follow as they migrate. Several methods have been used to trace the migratory pathways of neural crest cells. One approach is to employ vital labeling of premigratory neural crest cells, for example with lysinated rhodamine dextran (LRD) or the lipophilic dye, DiI, and subsequently following migrating cells in living embryos over time. Alternatively, immunohistochemical markers and molecular probes speciﬁc for neural crest cells often are used to assess the distributions of cells in ﬁxed tissues. In all species studied thus far, neural crest cells migrate along stereotyped routes. For reasons of space and simplicity, we focus on the migration of neural crest cells in the trunk. Discussions of cranial neural crest migration are available elsewhere (e.g., Köntges and Lumsden, 1996; Kulesa, 2004; Kulesa et al., 2004; Lumsden et al., 1991; Noden, 1991; Trainor et al., 2003). There are two principal migratory pathways for neural crest cells in the trunk (reviewed in Erickson, 2003; Erickson and Perris, 1993). One subpopulation of neural crest cells follows a ventromedial, or medial, pathway between the somite and neural tube. Glial and neurogenic precursors of the peripheral nervous system are among the neural crest cells that differentiate along this path, although in ﬁshes

and amphibians some of the neural crest cells in this path can also differentiate as pigment cells. A second subpopulation of neural crest cells migrates along a dorsolateral, or lateral, path between the somites and the overlying epidermis. Neural crest cells in this path give rise to pigment cells (reviewed in Erickson, 1993). In ﬁshes and amphibians, some neural crest cells take a third pathway and migrate dorsally into the developing ﬁn. In birds and mammals, neural crest cells in the dorsolateral path differentiate exclusively as pigment cells. Similarly in ectotherms, there is no deﬁnitive evidence that neural crest cells in the dorsolateral path contribute to any neural crest derivative other than pigment cells. Conversely, there is a bias toward nonpigment cell phenotypes in ventromedially migrating neural crest. This correlation between migratory path and differentiated phenotype is well documented, but its developmental basis is controversial. There is evidence not only to support the notion that environmental factors encountered within a particular migratory path dictate neural crest fate, but also conversely, that prior speciﬁcation dictates the choice of migratory path (discussed in Erickson, 2003; Le Douarin et al., 1993). The developmental potential of neural crest cells has been studied by a variety of means, including clonal analysis of neural crest differentiation and grafting neural crest cells or neural tube segments to ectopic axial levels and determining the differentiated phenotypes of the transplanted cells. Taken together, these studies suggest that most early neural crest cells are multipotent but become fate-restricted gradually over time (reviewed in Weston, 1991; Stemple and Anderson, 1993; Le Douarin et al., 1993, 2004; see below). Deﬁning the precise lineage segregation of pigment cells has been difﬁcult for two reasons. First, pigment cells differentiate from neural crest cells at most, if not all, axial levels of the trunk. As a result, heterotopic transplantation studies of the sort used to study the differentiation of other crest subpopulations in ectopic locations are not useful for studying the commitment of neural crest cells to a pigment cell fate. Second, markers for pigment cell precursors at the earliest stages of differentiation were not available until relatively recently. Consequently, early studies inferred that a particular cell was a pigment cell precursor simply by virtue of it being in the dorsolateral path but could not further evaluate the cell’s phenotype. Now, however, there is a substantial arsenal of lineage-speciﬁc markers for pigment cell precursors in most taxa. As detailed below, these markers have proven critical in studying the factors involved in pigment cell development.

Fishes Fishes provide three advantages for studies of pigmentation. First, the embryos of many species are oviparous and transparent so that neural crest cells and chromatophores can be observed throughout early development. Second, some ﬁshes have short generation times and can be bred and reared in the laboratory, permitting genetic approaches to studies of pigment cells and pigment pattern development. Third, 109

CHAPTER 5 Early larva

Adult

Mel

Mel

Xan Irid

Mel Irid

Mel

pigment patterns of ﬁshes are exceptionally diverse across taxa (e.g., Orton, 1953; Quigley et al., 2005; Riehl, 1991), allowing for comparative analyses of pigment cell development. To date, the advantages of ﬁsh are being exploited most fully in the tropical, freshwater zebraﬁsh Danio rerio, as well as the freshwater medaka Oryzias latipes. Several sources provide information on staging, husbandry, experimental techniques, as well as the histories of these species as research models [zebraﬁsh: Grunwald and Eisen (2002), Kimmel et al. (1995), Westerﬁeld (2000), Zebraﬁsh Information Network (http://zﬁn.org); medaka: Iwamatsu (2004), Shima and Mitani (2004), Medaka Homepage (http://biol1.bio.nagoya-u.ac. jp:8000/)]. Studies of these species have revealed numerous similarities in the genetic and cellular bases for neural crest and pigment cell development between teleosts and amniotes (see below). Recently several more extensive reviews of teleost pigment pattern formation have appeared (Kelsh, 2004; Parichy, 2003; Quigley and Parichy, 2002; Rawls et al., 2001). Like other vertebrates, a neural crest origin for ﬁsh pigment cells was long suspected (Borcea 1909), though not veriﬁed until considerably later (Humm and Young, 1956; Kajishima, 1958; Lopashov, 1944; for experiments on lampreys, see Newth, 1951, 1956). Early light microscopic studies of neural crest cells and chromatophores in teleosts identiﬁed these cells by location, shape, or externally visible pigmentation (e.g., Milos and Dingle, 1978a; Orton, 1953; Shephard, 1961). Grafting of [3H]thymidine-labeled cells also has been employed to examine neural crest migration (Lamers et al., 1981). Similarly, the HNK-1 antibody, which recognizes migrating neural crest cells of avian, rat, and turtle embryos (see below), has been used to identify neural crest cells in some 110

Fig. 5.1. Different pigment patterns of early larval and adult zebraﬁsh (see also Plate 5.1, pp. 494–495). Mel, melanophore stripes; Xan, xanthophores; Irid, iridophores.

species (Sadaghiani and Vielkind, 1989, 1990a; Sadaghiani et al., 1994). Finally, vital dyes have been used to label neural crest cells and their derivatives in zebraﬁsh (e.g., Dutton et al., 2001; Raible and Eisen, 1994; Smith et al., 1994; Vaglia and Hall, 2000). Teleost chromatophores retain their pigments intracellularly and pigment patterns thus reﬂect the numbers and arrangements of the various chromatophore classes (black melanophores, yellow xanthophores, iridescent iridophores, white leukophores, and red erythrophores). The precise complements of these chromatophores differ among species. For example, erythrophores are present in several Danio species, though not in zebraﬁsh (McClure, 1999; Quigley et al., 2004, 2005). Adding to the complexity of teleost pigment cell development, pigment patterns of many species differ between early larval and adult stages (Fig. 5.1). Early larval patterns of many species (including zebraﬁsh and medaka) comprise stripes of melanophores at the dorsal and ventral edges of the myotomes, along the horizontal myoseptum, and over the dorsal and ventral surfaces of the yolk (Kimmel et al., 1995; Kirschbaum, 1975; Lamoreux et al., 2005; Milos and Dingle, 1978a, b; Milos et al., 1983; Orton, 1953; Quigley et al., 2004). Iridophores or leukophores are found within the melanophore stripes, whereas xanthophores are widely dispersed outside of the stripes. This early larval pigment pattern arises during embryogenesis and is largely completed by hatching. After several days to weeks depending on the species, early larval pigment patterns are transformed into adult pigment patterns that are exceptionally diverse among species (Booth,

REGULATION OF MELANOBLAST MIGRATION AND DIFFERENTIATION

1990; Kirschbaum, 1975; Johnson et al., 1995; Mabee, 1995; Parichy and Johnson, 2001). For example, even within the genus Danio, different species exhibit horizontal stripes of varying number, width, and distinctiveness, vertical bars, uniform patterns, and more ornate patterns (Quigley et al., 2004, 2005). Pigment patterns are even more diverse among other groups including cichlids and coral reef ﬁshes (Allender et al., 2003; Lorenz, 1962; Streelman et al., 2003).

Morphogenesis of Fish Chromatophores Prospective neural crest cells in most vertebrates are found within the neural folds, which fuse to form a neural tube. In ﬁshes, the neural tube arises instead from a ventral thickening of the dorsomedial ectoderm termed the “neural keel.” Subsequently, the neural keel delaminates from surrounding epidermal ectoderm and a central canal develops to generate the deﬁnitive neural tube (Lowery and Sive, 2004; Schmitz et al., 1993). Neural crest cells ﬁrst become apparent along the dorsal surface of the developing neural tube as the neural keel separates from the epidermis (Lamers et al., 1981; Lowery and Sive, 2004; Raible et al., 1992; Sadaghiani and Vielkind, 1989, 1990a; Sadaghiani et al., 1994; Schmitz et al., 1993). The details of neural crest migration in ﬁshes are best documented in zebraﬁsh. After segregating from the neural keel, zebraﬁsh neural crest cells migrate along the two major pathways discussed previously: ventromedially between the somites and neural tube, and dorsolaterally between the somites and epidermis (Raible and Eisen, 1994; Raible et al., 1992; Vaglia and Hall 2000; Wood et al., 1992). In addition, some neural crest cells migrate dorsally and localize in the ﬁns. Zebraﬁsh neural crest cells in all pathways give rise to pigment cells (Raible and Eisen, 1994). The mechanisms underlying early neural crest cell dispersal and migration remain unknown in teleosts, though transient roles for somitic mesoderm in blocking early migration are likely (Jesuthasan, 1996). Some insights into early larval pigment cell morphogenesis have come from analyses of sparse mutant zebraﬁsh. In contrast to wild-type, sparse mutants have fewer melanophores and these cells are largely absent from the anterior head as well as ventral regions of the ﬂank. Molecular cloning identiﬁed sparse as a teleost ortholog of Kit (formerly c-kit; Parichy et al., 1999), long-studied for its role in mouse melanocyte development (see Mammals, below). kit encodes a type III receptor tyrosine kinase containing ﬁve extracellular immunoglobulin repeats and a split kinase domain. The ligand of Kit is Steel factor (SLF; also known as stem cell factor, mast cell growth factor, and kit ligand). Like other receptor tyrosine kinases, dimerization of Kit upon ligand binding is necessary for activation of the kinase domain (Blume-Jensen et al., 1991). Downstream signal transduction events initiated by Kit signaling in melanoblasts are largely unknown, but evidence suggests that the phospholipase C and phosphatidylinositol-3¢-kinase pathways are activated (reviewed in Galli et al., 1994). Zebraﬁsh kit is expressed by melanophores and their precursors. In null alleles, melanophores undergo apoptosis and are extruded from the

skin (Parichy et al., 1999). More recent analyses of an allelic series including conditional, temperature-sensitive alleles reveal genetically and temporally separable roles for kit in melanocyte migration (at early stages) and melanocyte survival (at later stages) (Rawls and Johnson, 2003; see also, Rawls and Johnson, 2000, 2001). Interestingly, the kit-related gene colony stimulating factor-1 receptor (csf1r; formerly, fms) is required for the survival and migration of zebraﬁsh xanthophores (Parichy et al., 2000b), suggesting that similar requirements for receptor tyrosine kinase activity are fulﬁlled by kit and csf1r in melanophore and xanthophore lineages, respectively. Recent studies have identiﬁed a second locus required for melanophore survival during early larval pigment pattern formation. Danio rerio touchtone mutants exhibit normal migration and speciﬁcation of melanophores, but melanoblasts and melanophores subsequently undergo apoptosis. Genetic mosaic analyses reveal that touchtone acts autonomously to the melanophore lineage (Arduini and Henion, 2004; Cornell et al., 2004). The touchtone mutant phenotype results from lesions in trpm7, which encodes a member of the transient receptor potential (TRP) family (Elizondo et al., 2005). The transmembrane trpm7 protein serves as both a cation channel and as a kinase (Wolf, 2004), and mutants have additional defects in kidney function and skeletal development. A role for trpm7 channel activity during melanophore development is demonstrated by the partial rescue of trpm7 mutant melanophores by simple calcium or magnesium supplementation to the medium. While trpm7 appears to act independently of the kit pathway during melanophore development (Cornell et al., 2004), the upstream regulators and downstream targets of trpm7 remain unknown. Beyond permissive roles for cell migration, survival, and proliferation, of particular interest are the instructive factors that generate speciﬁc arrangements of chromatophores. From several recent studies we now have a clearer notion of what these factors are both in early larval and adult pigment pattern formation. The arrangement of early larval melanophores depends in part on the stromal-derived factor 1a (sdf1a) chemokine. sdf1a is a chemoattractant that contributes to migration of primordial germ cells (PGCs) as well as the lateral lines, a bilateral sensory system that develops by the directed migration of a bullet-shaped “primordium” of cells within the epidermis. Both PGCs and lateral line primordia express cxcr4b, a Gprotein-coupled receptor for sdf1a, and migrate towards sources of sdf1a in the prospective gonads and along the horizontal myoseptum, respectively (David et al., 2002; Doitsidou et al., 2002; Li et al., 2004). A role for sdf1a in melanophore patterning is revealed by choker mutant zebraﬁsh (Kelsh et al., 1996). By early larval stages, these mutants have a “collar” of melanophores that covers the normally melanophore-free regions of the lateral myotomes. Myotome development is severely disrupted in choker mutants and sdf1a is expressed ectopically along the myotomes where melanophores accumulate. Morpholino knockdown of sdf1a 111

CHAPTER 5

in the choker mutant background reduces the numbers of collar melanophores, whereas ectopic sources of human sdf1a attract melanophores in wild-type embryos (V. Svetic and R. N. Kelsh, personal communication). During normal development then, sdf1a plausibly contributes to melanophore localization. Whether sdf1a acts directly on melanophores or through an intermediary signaling pathway remains unclear, particularly since melanophores do not express detectable levels of cxcr4b by in situ hybridization. Intriguingly, choker mutants lack xanthophores in the collar region. Genetic analyses show this defect is independent of the defect in melanophore patterning, whereas the defect in melanophore patterning is independent of the defect in xanthophore distribution. Development of the zebraﬁsh adult pigment pattern depends on a complex interplay between directed chromatophore migration and death, as well as differentiation (see below; Parichy and Turner, 2003b; Parichy et al., 2000b; Quigley et al., 2004, 2005). During metamorphosis, new xanthophores arise just ventral to the horizontal myoseptum and at the dorsal and ventral margins of the ﬂank, whereas new melanophores arise widely dispersed over the ﬂank and outside of the early larval stripes. Some of these “metamorphic” melanophores migrate short distances to join the ﬁrst two adult melanophore stripes, immediately dorsal and ventral to the horizontal myoseptum. Simultaneously, additional metamorphic melanophores develop already within the adult stripes. Many early larval melanophores initially within the lateral stripe along the horizontal myoseptum die, though some migrate to join the developing adult stripes, thereby producing a melanophore-free “interstripe region” that is, however, populated by xanthophores. As metamorphosis proceeds, the adult stripe borders become increasingly distinct. During juvenile and subsequent adult development, additional melanophore stripes are added dorsally and ventrally as the ﬁsh grows. In the mature adult pigment pattern of zebraﬁsh, dark stripes comprise melanophores and iridophores with occasional xanthophores, whereas light stripes comprise xanthophores and iridophores (Hirata et al., 2003; Parichy and Turner, 2003a). Dermal and epidermal chromatophores also are found outside of stripes; for example, scale melanophores contribute an overall dark cast to the dorsum of the ﬁsh. The morphogenetic bases for the zebraﬁsh adult striped pattern have been examined in several recent studies that have identiﬁed a critical role for interactions within and among chromatophore classes. The notion that chromatophore interactions could be important comes initially from the csf1r mutant phenotype (Parichy et al., 2000b; Fig. 5.2). As compared to wild-type, csf1r mutants have fewer adult melanophores and these cells fail to organize into normal stripes. Beyond the melanophore defect, csf1r mutants lack embryonic xanthophores (as mentioned above) as well as adult xanthophores. The correlated adult melanophore and xanthophore defects raised the possibility that interactions between these lineages might underlie melanophore stripe development. Support for this notion comes from two experimental paradigms (Parichy and Turner, 2003a). First, orga112

nized melanophore stripes are recovered in genetic mosaics in which melanophores and xanthophores are experimentally juxtaposed: when cells are transplanted between csf1r mutants (which lack xanthophores but retain melanophores) and nacre (mitfa) mutants [which lack melanophores but retain xanthophores (Fig. 5.2 and see below)] the resulting chimeras develop stripes. Second, if csf1r is conditionally activated or inactivated using a temperature-sensitive allele, xanthophores are gained or lost, respectively, and the number of xanthophores present correlates with the degree of melanophore stripe organization. Indeed, xanthophores and melanophore stripes can be gained or lost in a csf1r-dependent manner throughout development, revealing roles for csf1r in both initiating and maintaining stripe formation. Since csf1r is not detectably expressed by melanophores but is expressed by the xanthophore lineage, these results suggest that interactions between melanophores and csf1r-dependent cells of the xanthophore lineage are required for stripe organization. Chromatophore interactions probably also contribute to development of stripes on the ﬁns (Goodrich and Greene, 1959; Goodrich and Nichols, 1931; Goodrich et al., 1954; Parichy and Turner, 2003a). Analyses of other Danio species suggest that evolutionary changes in chromatophore interactions may underlie interspeciﬁc variation in adult pigment patterns (Parichy and Johnson, 2001; Quigley et al., 2005). Further indications that chromatophore interactions underlie stripe development come from studies of obelix (jaguar) and leopard mutant zebraﬁsh (Haffter et al., 1996; Johnson et al., 1995; Fig. 5.2). obelix mutants exhibit fewer and broader stripes than wild-type and xanthophores are intermingled with melanophores in the melanophore stripes (Maderspacher and Nüsslein-Volhard, 2003). leopard mutants often develop spots, but alleles of differing severity range in phenotype from irregular stripes to nearly uniformly distributed, intermingled melanophores and xanthophores (Asai et al., 1999; Maderspacher and Nüsslein-Volhard, 2003). Thus, mutant phenotypes may be interpretable in the context of defective boundary formation between chromatophore classes. Cell transplantation and genetic analyses indicate that obelix acts within melanophores to promote homotypic interactions between these cells, whereas leopard acts within melanophores and xanthophores to promote both homotypic and heterotypic interactions (Maderspacher and NüssleinVolhard, 2003). The obelix mutant phenotype results from mutations in kir7.1, encoding an inwardly rectifying potassium channel (S. L. Johnson and S. Kondo, personal communication); the cellular basis for translating channel activity into cellular pattern remains to be elucidated. leopard mutant phenotypes result from mutations in the gap junction gene, connexin-40, suggesting that direct cell–cell contacts and cytoplasmic communication may underlie leopard-dependent chromatophore interactions and boundary formation (S.L. Johnson, personal communication). Interestingly, N-CAM and N-cadherin had previously been associated with xanthophores but not other chromatophore classes in O. latipes (Fukuzawa and Obika, 1995), consistent with roles for dif-

REGULATION OF MELANOBLAST MIGRATION AND DIFFERENTIATION

Wild-type

picasso (erbb3)

puma

panther (csf1r)

sparse (kit)

panther (csf1r); sparse (kit)

rose (ednrb1)

panther (csf1r); rose (ednrb1)

sparse (kit); rose (ednrb1)

nacre (mitfa)

leopard (cx40)

jaguar (kir7.1)

Fig. 5.2. Mutational analyses allow a genetic dissection of adult pigment pattern development in zebraﬁsh. Shown are wild-type zebraﬁsh as well as single and double mutant zebraﬁsh with corresponding pigment pattern defects (see also Plate 5.2, pp. 494–495). See text for details.

ferential adhesion in the sorting-out of speciﬁc chromatophore classes, although this model has yet to be tested directly. Besides the spatial arrangements of chromatophore classes across the ﬂank, different chromatophores types occupy different strata within the skin itself (Hawkes, 1974a, b). In zebraﬁsh, light stripes comprise xanthophores superﬁcial to iridophores, whereas dark stripes comprise occasional xanthophores superﬁcially, followed by iridophores, then melanophores, and additional iridophores more internally. Moreover, the organization of iridophore reﬂecting platelets differs depending on whether or not these cells are associated with xanthophores or melanophores (Hirata et al., 2003). The molecular signals associated with these arrangements are not known.

Differentiation of Fish Chromatophores Factors controlling the speciﬁcation and differentiation of teleost chromatophores are beginning to be elucidated. An early step in neural crest speciﬁcation towards chromatophore fates requires the transcription factor sox10, as revealed by analyses of colourless mutant zebraﬁsh in which the sox10 gene is disrupted (Dutton et al., 2001; Kelsh and Eisen, 2000). sox10 (colourless) mutants have severe defects in chromatophores, glia, and neurons: precursors to these lineages fail

to leave the neural crest and instead undergo apoptosis. In contrast, ectomesenchymal neural crest derivatives (e.g., craniofacial skeleton and ﬁn mesenchyme) are unaffected. Nonectomesenchymal sox10-dependent neural crest cells often are fate-restricted even prior to leaving the neural crest, as revealed by elegant single cell lineage analyses (Dutton et al., 2001; Raible and Eisen, 1994). Such analyses do not yet provide a clear picture of lineage relationships between chromatophores and other nonectomesenchymal lineages, or even among chromatophore classes themselves. (For a fuller discussion of neural crest speciﬁcation, see Birds, below.) Nevertheless considerable progress has been made in deﬁning the mechanisms of fate speciﬁcation and differentiation in the melanophore lineage. An important advance came from the isolation of nacre mutant zebraﬁsh that lack melanophores. This mutant phenotype results from mutations in microphthalmia-a (mitfa) (Lister et al., 1999), encoding a member of the basic-loophelix-leucine-zipper (bLH-ZIP) family of transcription factors (Hodgkinson et al., 1993; see Steingrimsson et al., 2004 for a recent review). Mammalian orthologs of mitfa have long been studied and are known to directly upregulate genes required for melanin synthesis (see Mammals, below). Additional studies of zebraﬁsh have provided insights into how 113

CHAPTER 5

mitfa is itself regulated, with important roles identiﬁed for both intrinsic and extrinsic factors. An intrinsic factor relative to the melanophore lineage is the sox10 transcription factor. mitfa expression is curtailed in sox10 (colourless) mutants (Dutton et al., 2001). The sox10-dependence of melanophore development lies entirely in sox10-dependent regulation of mitfa: sox10 binds and activates the mitfa promoter, sox10 overexpression upregulates mitfa expression in vivo, and forced mitfa expression rescues melanophore development in sox10 mutant backgrounds (Elworthy et al., 2003). An extrinsic factor relative to the melanophore lineage lies in the Wnt signaling pathway. Wnt1 and Wnt3a are expressed in the dorsal neural keel. This expression domain has functional signiﬁcance in promoting chromatophore over nonchromatophore fates as revealed by injections of single premigratory neural crest cells with activating or inhibitory mRNAs for the Wnt signal transduction pathway (Dorsky et al., 1998). This effect is mediated, at least in part, by the binding of Wnt-responsive Lef1/T-cell factor (TCF) transcription factors within the mitfa promoter, which upregulate mitfa transcription and thus melanophore fate speciﬁcation (Dorsky et al., 2000a, b). More recent studies using a heat shock inducible transgene for a dominant negative form of TCF demonstrate that Wnt signals are required at several distinct steps in the neural crest–melanophore lineage (Lewis et al., 2004; see also Birds, below). Beyond the critical roles for mitfa, sox10, and Wnts in melanophore speciﬁcation, a large number of mutants have been isolated in zebraﬁsh and medaka that identify genes likely to function primarily in melanophore differentiation (Kelsh et al., 1996, 2000, 2004). Many of these probably affect melanin synthesis or melanosome biogenesis (e.g., Fukamachi et al., 2001; Schonthaler et al., 2005). One gene likely to regulate these processes encodes transcription factor activating protein-2a (tfap2a). The zebraﬁsh lockjaw and mont blanc mutant phenotypes result from mutations in tfap2a and have severe defects in melanophore as well as iridophore development, in addition to defects in craniofacial, glial, and neural derivatives of the neural crest (Barrallo-Gimeno et al., 2004; Knight et al., 2003, 2004). The failure of melanophore development results at least in part from a tfap2a-dependence of kit expression, and epistasis analyses combined with morpholino knockdown support a model in which tfap2a and kit act in a common pathway (O’Brien et al., 2004). As melanoblasts express early markers of the melanophore lineage, and some melanophores develop but are lightly pigmented, it appears tfap2a allows the terminal differentiation of this cell type. While considerable effort is directed towards understanding speciﬁcation and differentiation of chromatophores during embryonic stages, mutational analyses have also started to reveal genetic requirements of metamorphic melanophores that contribute to adult pigment patterns. For example, puma mutant zebraﬁsh have a severe deﬁcit of metamorphic melanophores despite having a normal complement of early larval melanophores (Parichy and Turner, 2003b; Parichy 114

et al., 2003; Fig. 5.2). puma is required during metamorphosis for promoting melanoblast and melanophore development and acts autonomously to the melanophore lineage. A superﬁcially similar phenotype is observed for picasso mutants (Quigley et al., 2004; Fig. 5.2), which result from mutations in the receptor tyrosine kinase gene erbb3 (Lyons et al., 2005; E. Budi and D. Parichy, unpublished data). Nevertheless, it has yet to be determined whether puma and picasso (erbb3) function primarily to promote metamorphic melanophore speciﬁcation and differentiation, or survival and proliferation. Regardless of the precise mechanism, comparative analyses demonstrate that most adult pigment patterns among danios arise from metamorphic melanophores that differentiate from latent precursors during the larval-to-adult transformation (Quigley et al., 2004); understanding the factors that promote the establishment, maintenance, and recruitment of such latent precursors is thus critical to elucidating the mechanisms underlying naturally occurring pigment pattern variation in this group. As an extreme example of this notion, the zebraﬁsh sister species, D. nigrofasciatus, mostly lacks metamorphic melanophores and their precursors owing to evolutionary changes that are extrinsic to this melanophore lineage and likely to involve the puma pathway (Quigley et al., 2004). Despite their apparent uniformity, metamorphic melanophores actually comprise two genetically and temporally distinct populations, as revealed by analyses of kit, csf1r, and endothelin receptor b1 (ednrb1) mutants (Johnson et al., 1995; Parichy et al., 1999, 2000a, b). kit mutants not only lose their early larval melanophores, but lack early-appearing, dispersed metamorphic melanophores; nevertheless, these mutants ultimately recover metamorphic melanophores, already situated in stripes, during late metamorphosis and juvenile development. In contrast, csf1r and ednrb1 mutants have normal numbers of early-appearing, initially dispersed metamorphic melanophores and instead are deﬁcient for late-appearing metamorphic melanophores at sites of stripe formation. That early kit-dependent and late csf1r- and ednrb1-dependent melanophores are distinct populations is supported by epistasis analyses: in contrast to single mutants, ﬁsh doubly mutant for kit and csf1r, or kit and ednrb1, lack nearly all body melanophores (Johnson et al., 1995; Parichy et al., 2000b; Fig. 5.2). Additional discussion of endothelin receptors is provided in sections on Birds and Mammals, below. In contrast to melanophores, relatively less is known about the speciﬁcation and differentiation of xanthophores, iridophores, or other chromatophore classes. Early larval iridophores are reduced in tfap2 mutants whereas adult iridophores are absent in ednrb1 mutants. Molecular identiﬁcation of additional mutants affecting iridophore development [e.g., shady and parade (Kelsh et al., 1996)] will surely provide valuable new insights. Virtually nothing is known about xanthophore fate speciﬁcation, though precursors of these cells and xanthophore numbers are somewhat reduced in mitfa mutants (Lister et al., 1999; Parichy et al., 2000b). Xanthophore pigments are

REGULATION OF MELANOBLAST MIGRATION AND DIFFERENTIATION

derived, in part, from the pteridine pathway (Ziegler, 2003; Ziegler et al., 2000) and enzymes within this pathway are expressed by xanthophore precursors, although also in some other lineages, including melanophores (Parichy et al., 2000b; Parichy, unpublished data), suggesting caution in their use as lineage markers. Mutational analyses reveal that xanthophore differentiation requires PAM (protein associated with cmyc), which regulates pteridine synthesis enzymes in a cell autonomous manner (Le Guyader et al., 2005), as well as somatolactin, which acts as an endocrine factor (Fukamachi et al., 2004).

Mutants Besides those discussed above, a very large number of other mutants affecting chromatophore development have been isolated, particularly in zebraﬁsh (Golling et al., 2002; Haffter et al., 1996; Kelsh et al., 1996; Odenthal et al., 1996; Rawls et al., 1993) and O. latipes (Kelsh et al., 2004). These mutants, high-throughput screens for chemically-induced phenotypes (Peterson et al., 2000), and genomic resources (Pickart et al., 2004) offer outstanding opportunities for dissecting morphogenesis and differentiation, as well as the evolution of these processes. Descriptions of mutant screens and on-line resources for these mutants can be found at: ∑ http://zﬁn.org ∑ http://biol1.bio.nagoya-u.ac.jp:8000/ ∑ http://protist.biology.washington.edu/dparichy/ ImagesMutants.htm

Amphibians Amphibians offer several advantages for studies of pigment cells and pigment pattern formation. For example, embryos of many species are quite large and develop oviparously. Moreover, development rate often can be controlled by incubating embryos at a wide range of temperatures. This makes amphibian embryos highly accessible to both observation and experimentation. Finally, amphibians offer a wide range of patterns to study: not only do different taxa exhibit an extraordinary diversity of larval and postmetamorphic patterns [e.g., urodeles (Bishop, 1941; Epperlein and Löfberg, 1990; Olsson, 1993, 1994; Pﬁngsten and Downs, 1989; Twitty, 1936, 1966), reviewed in Parichy, 1996b; anurans (Altig and Johnston, 1989; Andres, 1963; Bagnara, 1982; Elias, 1941; Ellinger, 1979; Frost and Robinson, 1984; Gosner and Black, 1957; Smith-Gill, 1974), reviewed in Frost-Mason et al. (1995)], but mutations have been identiﬁed in some species that speciﬁcally affect chromatophores or pigment pattern formation (see below). Salamander embryos were used by DuShane (1934, 1935, 1938) in his pioneering studies that ﬁrst revealed a neural crest origin for chromatophores. These ﬁndings were later extended to anurans (Stevens, 1954) and inspired numerous analyses of pigment pattern development in amphibians. Several reviews provide an introduction to these classical studies (Bagnara, 1983; DuShane, 1943; Epperlein and Löfberg, 1993; Erickson, 1993; Frost and Malacinski, 1980; Hall and

Hörstadius, 1988; Lehman and Youngs, 1959; Parichy, 1996b; Rawles, 1948; Twitty, 1949, 1953, 1966). Amphibian chromatophores in the integument can localize in both the dermis and the epidermis, though dermal chromatophores are primarily responsible for distinctive pigment patterns in most taxa (e.g., Bagnara et al., 1968; DuShane, 1943; Stearner, 1946).

Morphogenesis of Amphibian Chromatophores In salamanders, rounded, presumptive neural crest cells are ﬁrst detectable within the neural folds (Hirano and Shirai, 1984; Raven, 1931; Spieth and Keller, 1984). As the neural folds fuse at the midline, prospective neural crest cells form a wedge within the neuroepithelium and ﬁnally emerge as a distinctive but transient cord or “crest” running anteroposteriorly along the dorsal aspect of the completed neural tube (Fig. 5.3). Presumptive neural crest cells also have been identiﬁed in the neural folds of the anuran Xenopus laevis (Mayor et al., 1995; Sadaghiani and Thiébaud, 1987; Schroeder, 1970). Many techniques have been used to identify neural crest cells in amphibian embryos, including light and electron microscopy (Hirano and Shirai, 1984; Löfberg and Ahlfors, 1978; Löfberg et al., 1980, 1985; Spieth and Keller, 1984), vital labeling (Collazo et al., 1993; Detwiler, 1937), and intrinsic characteristics or labeling of transplanted cells in chimeric embryos (Chibon, 1967; Krotoski et al., 1988; MacMillan, 1976; Moury and Jacobson, 1989, 1990; Sadaghiani and Thiébaud, 1987; Raven, 1931, 1936). Finally, amphibian — like teleost — chromatophores often produce pigment prior to reaching their ﬁnal destination and can be observed directly (Epperlein and Claviez, 1982; Epperlein and Löfberg, 1990; Keller and Spieth, 1984; Tucker and Erickson, 1986a; Twitty, 1945). Amphibian neural crest cells disperse along dorsolateral and ventromedial pathways and also are found within the expanding ﬁn. In salamanders, most or all neural crest cells that contribute to pigment patterns on the dorsal ﬂank of young larvae migrate dorsolaterally between the somites and epidermis (Epperlein and Löfberg, 1990; Olsson and Löfberg, 1992). Neural crest cells traveling dorsolaterally are probably the major contributors to externally visible pigment patterns in most frog tadpoles as well (Andres, 1963; Tucker, 1986). In contrast, most X. laevis neural crest cells and melanophores fail to migrate subepidermally in the anterior trunk and instead remain near the dorsal apex of the myotomes or ﬁrst travel medially and only secondarily migrate to the epidermis (Andres, 1963; Collazo et al., 1993; Krotoski et al., 1988; MacMillan, 1976; Tucker, 1986). Medial migration of melanophores or their precursors in other amphibians is suggested by the observation that many amphibians exhibit chromatophores lining internal structures (DuShane, 1943; Tucker and Erickson, 1986a). The factors controlling amphibian neural crest and chromatophore morphogenesis are not well understood, but results from many studies suggest roles for the extracellular matrix (ECM). Work in Ambystoma mexicanum suggests that changes in the composition of the ECM may inﬂuence the 115

CHAPTER 5

Fig. 5.3. Neural crest cells (arrows) emerging from the neural tube of a salamander embryo. Bar = 200 mm.

initial dispersal and migration of neural crest cells (Löfberg and Ahlfors, 1978; Löfberg et al., 1980, 1985). A variety of ECM components have been identiﬁed in the vicinity of early or later migrating amphibian neural crest cells and chromatophores, including ﬁbronectin, laminin, collagens type I and III, chondroitin and keratan sulfate proteoglycans, tenascin, hyaluronan, and molecules that bind the lectin peanut agglutinin (PNA) (Epperlein and Löfberg, 1990; Epperlein et al., 1988; Olsson et al., 1996; Parichy, 1996a, 2001a; Perris et al., 1990; Tucker, 1986; Tucker and Erickson, 1986a, b). Several studies have examined the behavior of salamander neural crest cells on ECM substrata in vitro (Epperlein et al., 1988; Perris and Johansson, 1987, 1990; Tucker and Erickson, 1986a, b). These investigations have shown that salamander neural crest cells can disperse and migrate on a variety of ECM components, including ﬁbronectin, vitronectin, laminin, and collagens type I, IV, and VI. In contrast, puriﬁed tenascin inhibits the adhesion of neural crest cells, and motility of these cells can be modulated in a concentrationdependent manner by addition of chondroitin sulfate proteoglycans and hyaluronan. In the early larvae of several salamander taxa, melanophores ﬁrst are uniformly distributed over the ﬂank but subsequently concentrate near the dorsal apex of the myotomes and along the dorsal margin of the yolk mass. Because xanthophores typically are present over the lateral face of the myotomes, the resulting pattern consists of dark, horizontal dorsal and lateral stripes bordering a pale “melanophore-free region” along the middle of the ﬂank (Parichy, 1996b, 2001a, b). In one of these taxa, Taricha torosa, ﬁbronectin is abundant at sites of melanophore localization, whereas hyaluronan is abundant within the dorsal ﬁn matrix where melanophores are not found (Tucker and Erickson, 1986a, b). In Triturus alpestris, the local appearance of tenascin and chondroitin sulfate proteoglycans over the lateral face of the myotomes correlates with the evacuation of 116

melanophores from this region (Epperlein and Löfberg, 1990). Moreover, interspeciﬁc differences in tenascin and laminin correlate with species-speciﬁc melanophore distributions (Parichy, 2001a). In several other taxa within the families Ambystomatidae and Salamandridae, the lateral line sensory system (Northcutt, 1992; Northcutt et al., 1994) is directly responsible for excluding melanophores from the middle of the ﬂank (Parichy, 1996a, c). Lateral line-dependent perturbations of the subepidermal basement membrane and increased deposition of tenascin correlate with the evacuation of melanophores from this region (Parichy 1996a, 2001a). Changes in the ECM also may inﬂuence chromatophores and chromatoblasts during later development. In A. t. tigrinum for example, staining for laminin, tenascin, and PNA-binding activity increase dramatically during terminal formation of the early larval pigment pattern, probably reﬂecting the elaboration of a largely acellular dermis that is hypothesized to stabilize melanophore positions (Parichy, 1996c; Stearner, 1946). Likewise, breaks in the collagenous basement membrane and the appearance of ﬁbronectin “tracts” correlate with changes in melanophore positions at metamorphosis in the frog Rana esculenta (Denèﬂe and Lechaire, 1990, 1991, 1992; Denèﬂe et al., 1987). Another class of factors that probably inﬂuences the morphogenetic behavior of chromatophores is interactions among the cells themselves, as in teleosts. Based on a series of studies in vitro, Twitty (1944) and Twitty and Niu (1948) suggested that the directed dispersal of neural crest cells and melanophores results from the production of diffusible substances that repel neighboring cells. Although such negative chemotaxis has not been ruled out in urodeles, attempts to verify these conclusions using quail neural crest cells have not been successful (Erickson and Olivier, 1983). More recently, melanophore and neural crest cell dispersal has been suggested to result from contact inhibition of movement (Abercrombie, 1970) coupled with high cell densities in dorsal regions of the

REGULATION OF MELANOBLAST MIGRATION AND DIFFERENTIATION

ﬂank [Tucker and Erickson (1986a); but see Keller and Spieth (1984)], concordant with similar proposals in avian embryos (Rovasio et al., 1983; Erickson, 1985). In contrast to repellent interactions that appear to mediate pigment cell dispersion, adhesive interactions among melanophores have been suggested to contribute to the condensation of these cells during stripe formation in T. torosa (Parichy, 1996a; Twitty, 1945). Cell–cell interactions also may be involved in the development of vertical barring patterns in several salamanders within the family Ambystomatidae (Epperlein and Löfberg, 1990; Lehman, 1954, 1957; Olsson, 1994; Olsson and Löfberg, 1992; Parichy, 1996a, b, 2001b); similar patterns also are present in some frog tadpoles (Altig and Johnston, 1989; Gosner and Black, 1957). In these taxa, melanophores scatter widely over the somites during early stages while xanthophores are retained in aggregates at variable intervals dorsal to the neural tube. Xanthophores subsequently disperse from these aggregates, and melanophores on the ﬂank recede short distances from the advancing xanthophores. This results in alternating vertical bars of melanophores and xanthophores in young larvae. Epperlein and Löfberg (1990) hypothesize that aggregates arise at the level of the premigratory neural crest by xanthophores “sorting out” from surrounding melanophores via differential adhesion (Steinberg, 1970). Observations from experimentally manipulated and mutant embryos are consistent with this idea. Nevertheless, the events of aggregate formation have yet to be described in vivo. During later stages, repulsive interactions between melanophores and xanthophores appear to drive morphogenesis of the deﬁnitive vertical barring pattern (Epperlein and Löfberg, 1990) though the underlying mechanisms have yet to be established. In addition to interactions among chromatophores and between chromatophores and the ECM, gross physical features of the extracellular environment probably inﬂuence the migration and localization of amphibian chromatophores. For example, a physical barrier near the dorsal margin of the yolk mass may contribute to the failure of T. torosa melanophores to migrate ventrally over the lateral plate mesoderm (Twitty, 1936). Melanophores of this species also localize along the lateral line nerve perhaps due to contact guidance (Dunn, 1982; Tucker and Erickson, 1986a). Finally, the lateral line nerve in A. tigrinum is hypothesized to act as a physical barrier that prevents dorsal melanophores from migrating further ventrally (Parichy, 1996c). A variety of hormones and growth factors also could inﬂuence the behavior of embryonic amphibian neural crest and pigment cells. Candidate molecules have been identiﬁed in teleost and amniote embryos, but the distributions and functions of these molecules and their receptors are only beginning to be investigated in amphibians at stages relevant to early pigment pattern formation (e.g., Baker et al., 1995). Finally, externally visible pigment patterns of most amphibians change markedly during later larval stages and especially during and after metamorphosis (Bagnara, 1982; DuShane,

1943; Fernandez and Collins, 1988; Lehman, 1953; Lehman and Youngs, 1959; Niu and Twitty, 1950; Parichy 1998, 2001b; Smith-Gill, 1974; Stearner, 1946; Twitty, 1936). These changes presumably reﬂect persistent or renewed migration of existing pigment cells as well as neural crest cells or chromatoblasts that have not yet overtly differentiated. Nevertheless, it seems likely that these chromatophore movements are inﬂuenced by endocrine-derived hormones (reviewed in FrostMason et al., 1995). Hormones and localized growth factors probably also inﬂuence the massive restructuring of skin that occurs during metamorphosis (Heatwole and Barthalmus, 1995), which may indirectly impact the establishment of a new pigment pattern. For example, thyroid hormones trigger a cascade of events during metamorphosis, including extensive remodeling of the ECM, cell death, proliferation, and differentiation (e.g., Patterton et al., 1995). Intriguingly, thyroxine stimulates the migration of melanophores from the dermis to the epidermis in cultured frog integument, possibly reﬂecting a direct effect of the hormone on melanophores (Yasutomi, 1987).

Differentiation of Amphibian Chromatophores At least some amphibian neural crest cells begin to differentiate into melanophores and xanthophores shortly after dispersing, or even while in the premigratory position dorsal to the neural tube (Epperlein and Löfberg, 1984, 1990; Olsson and Löfberg, 1992; Tucker and Erickson, 1986a). In contrast, iridophores typically are detectable only after the appearance of melanophores and xanthophores. It seems likely that additional populations of chromatoblasts remain covert within the integument and differentiate at later larval stages, during metamorphosis, or during adult life (Bagnara et al., 1979; Brown, 1997; DuShane, 1943; Fernandez and Collins, 1988; Frost et al., 1984a; Ohsugi and Ide, 1983; Parichy, 1998, 2001b; Stearner, 1946). In A. mexicanum, differentiation into melanophores and xanthophores is stimulated by serum (Dean and Frost-Mason, 1991), or alternatively if the cells are co-cultured with neural tube, epidermis, or ECM isolated from the dorsolateral (but not medial) pathway (Perris and Löfberg, 1986; Perris et al., 1988). Similar requirements for serum are observed in cultures of neural crest cells isolated from T. torosa (Tucker and Erickson, 1986b; Twitty, 1944, 1945) and X. laevis (Fukuzawa and Ide, 1988). One molecule known to stimulate melanogenesis speciﬁcally is a-melanocyte stimulating hormone (a-MSH), which is produced in the pituitary gland of adult animals. a-MSH originally was identiﬁed as a circulating hormone that causes the dispersion of melanin-containing granules (melanosomes) within melanocytes (e.g., Bagnara and Hadley, 1969; Fernandez and Bagnara, 1991; Sawyer et al., 1983). In addition to these short-term physiological responses, a-MSH probably also mediates long-term morphologic changes in pigmentation (reviewed in Bagnara and Hadley, 1969). In vitro, a-MSH stimulates both proliferation and melanization of anuran neural crest cells (Fukuzawa and Bagnara, 1989; 117

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Wilson and Milos, 1987) and melanophores (Fukuzawa and Bagnara, 1989). A possible role for a-MSH or related peptides during early development also has been suggested in a preliminary report demonstrating a-MSH immunoreactivity in the epidermis of A. mexicanum embryos and larvae (FrostMason et al., 1992). It will be interesting to see if a-MSH activity contributes to the early differentiation of neural crest cells into chromatophores, particularly in light of taxon- and stage-speciﬁc differences in the ability of epidermis to promote melanophore differentiation (DeLanney, 1941; Thibaudeau and Frost-Mason, 1992; Twitty, 1936). In addition to a-MSH, presently unidentiﬁed factors in the dorsal integument and serum of Rana pipiens and X. laevis stimulate both melanization and outgrowth in cultured X. laevis neural crest cells (Fukuzawa and Bagnara, 1989; Mangano et al., 1992). Conversely, a partially puriﬁed factor from ventral integument of Rana and X. laevis inhibits outgrowth and melanization of these cells (Fukuzawa and Bagnara, 1989; Fukuzawa and Ide, 1988), whereas medium containing factors derived from ventral integument stimulates adhesion and proliferation of iridophores in vitro (Bagnara and Fukuzawa, 1990). An antibody raised against a putative “melanization-inhibiting factor” has been shown to block inhibitory effects in vitro, and this antibody stains the ventral but not dorsal integument of R. pipiens (Fukuzawa et al., 1995). Further characterization of this factor could provide important insights into the control of regional differences in pigmentation (e.g., the dark dorsal and white bellies seen in many amphibians and other vertebrates).

Mutants Numerous pigment mutants have been found across several amphibian taxa, including R. pipiens (Davison, 1964; SmithGill, 1973, 1974, 1975; Volpe and Dasgupta, 1962), Bombina orientalis (Ellinger, 1980; Frost et al., 1982), X. laevis (Droin, 1992), and P. waltl (Collenot et al., 1989). Perhaps the most studied are those of A. mexicanum: melanoid, albino, axanthic, and white (reviewed in Frost and Malacinski, 1980; Frost et al., 1984b). Early larval “wild-type” A. mexicanum display abundant melanophores and xanthophores, whereas older larvae and paedomorphic adults exhibit melanophores, xanthophores, and iridophores in an irregular mottled green and black pattern that covers the integument. melanoid (m) In contrast to wild-type A. mexicanum, m/m animals exhibit few xanthophores as larvae or adults and completely lack iridophores at all stages of development (Frost et al., 1984c). Thus, melanoid adults are a velvety black. Genetic mapping places the melanoid locus on linkage group 14 (Smith et al., 2005). The melanoid defect is believed to act autonomously within the neural crest lineage (Sawada and Dalton, 1979). Total pigment cell numbers do not differ between melanoid and wild-type animals (Hoerter, 1977). Biochemically, the deﬁcit of xanthophores and iridophores correlates with a decrease in overall pteridine levels during 118

development (Frost et al., 1984c) as well as lower levels of pteridines compared to wild-type animals (Benjamin, 1970; Frost et al., 1984b; Thorteinsdottir and Frost, 1986). Treatment of wild-type A. mexicanum with an inhibitor of the pteridine synthesis enzyme xanthine dehydrogenase results in a phenocopy of the melanoid defect (Frost et al., 1989; Thorteinsdottir and Frost, 1986). albino (a) The a gene was originally introduced into A. mexicanum from the closely related A. tigrinum [an undertaking that required truly heroic efforts; see Humphrey (1967)]. Homozygotes for the albino gene completely lack melanin but do have unmelanized “melanophores” containing organelles typical of early stages in melanogenesis (Frost et al., 1986a). Because a/a individuals possess both xanthophores and iridophores, their overall coloration is a golden yellow. Albino A. mexicanum also exhibit differences in pteridine proﬁles within the integument compared to wild-type A. mexicanum (Frost et al., 1986a). The biochemical defect underlying the albino phenotype remains unknown. axanthic (ax) Little is known about the axanthic defect. The ax/ax A. mexicanum completely lacks iridophores whereas “xanthophores” are identiﬁable but remain unpigmented; melanophores are apparently unaffected (Frost et al., 1984a, 1986b). white (d) Homozygous d/d individuals lack all three types of chromatophores in the integument during most or all of their lives (Frost et al., 1984a), resulting in a white phenotype. Notably, eye pigmentation is unaffected, suggesting that the defect is speciﬁc for neural crest-derived pigment cells. Deﬁcient cutaneous pigmentation is thought to result from a failure of neural crest cells and chromatophores to disperse into the dorsolateral migratory pathway (Dalton, 1950; Keller et al., 1982), though lineage tracers have yet to be employed to test explicitly the relative roles of inhibited dispersal versus inhibited differentiation or survival [see also DuShane (1939)]. Indeed, some chromatophore precursors must disperse, because some d/d animals repigment spontaneously in haphazard patches after reaching sexual maturity (Frost and Malacinski, 1980; Frost et al., 1984b). The defect associated with the d gene traditionally has been attributed to a change within the epidermis, since wild-type epidermis grafted to white embryos permits melanophore dispersal whereas the converse does not (Keller et al., 1982). Löfberg et al. (1989) obtained analogous results when microcarriers coated with only white or wild-type subepidermal ECM were grafted between embryos. In contrast, microcarriers coated with subepidermal ECM from older white embryos permitted neural crest dispersal in both white and wild-type hosts. This led to the hypothesis that changes in the subepidermal ECM normally required for neural crest cell migration are transiently retarded in white embryos. Unfortunately, com-

REGULATION OF MELANOBLAST MIGRATION AND DIFFERENTIATION

parisons of white and wild-type ECM have yielded conﬂicting results (Epperlein and Löfberg, 1993; Olsson et al., 1996; Perris et al., 1990) and do not adequately explain what the white defect might be. Genetic mapping places d on linkage group 1, allowing the exclusion of several candidate genes, including steel, encoding the kit ligand, SLF (Parichy et al., 1999; Smith et al. 2005; Voss et al., 2001).

Reptiles Little is known concerning neural crest and pigment cell development in reptilian embryos. Hou and Takeuchi (1994) used the HNK-1 antibody to show that neural crest cells of the softshell turtle Trionyx sinensis migrate along both ventromedial and dorsolateral pathways, as in other vertebrates. Melanophores in this species are found cutaneously and internally (Hou and Takeuchi, 1991), suggesting that both ventromedially and dorsolaterally migrating neural crest cells can differentiate as melanophores. Intriguingly, early migrating neural crest cells also enter the myotome (Hou and Takeuchi, 1994). This could reﬂect enhanced invasiveness of a melanogenic subpopulation of neural crest cells traveling along the medial pathway, as has been suggested in the Silkie chicken (Faraco et al., 2001; Reedy et al., 1998b).

Birds Avian embryos are amenable to experimental manipulation at virtually all stages of embryogenesis and thus have been widely exploited as a model system for studying neural crest and pigment cell development. By far the two most commonly used avian species for these studies are the domestic chicken, Gallus gallus, and the Japanese quail, Coturnix japonica. Many studies have taken advantage of the fact that quail neural crest cells, when grafted into a chick embryo host, can be distin-

guished by their characteristic nuclear staining pattern with Feulgen dye (Le Douarin, 1982) or the quail-speciﬁc antibody QCPN. In addition, avian neural crest cells can be identiﬁed on the basis of their immunoreactivity with the HNK-1 antibody (Tucker et al., 1988) and by their expression of speciﬁc cadherins (Nakagawa and Takeichi, 1995) or transcription factors, such as Sox 10 (Cheng et al., 2000). The neural crest origin of avian melanocytes, the sole skin pigment cell type found in birds, was demonstrated over 50 years ago using experimental embryological techniques (Dorris, 1936, 1938, 1939; Ris, 1941; Watterson, 1942; Willier and Rawles, 1940). In adult birds, melanocytes are distributed in both the dermis and the epidermis (Hulley et al., 1991). In addition, melanocytes are found internally in some birds, for example the Silkie chicken (e.g., Faraco et al., 2001; Makita and Mochizuki, 1984; Makita and Tsuzuki, 1986).

Morphogenesis of Avian Melanocytes The dynamics of neural crest migration in the avian trunk have been well documented. In chickens, neural crest cells ﬁrst detach from the neuroepithelium in the trunk at stage 11 (Tosney, 1978) and invade the ventromedial pathway shortly thereafter (Bronner-Fraser, 1986; Loring and Erickson, 1987). These neural crest cells differentiate into the neurons and glial cells of the peripheral nervous system. The entry of avian neural crest cells into the dorsolateral path is delayed by about 24 hours with respect to ventromedial migration and begins at stage 20 at the level of the forelimb (Erickson et al., 1992; Serbedzija et al., 1989). Neural crest cells destined for the dorsolateral path initially pause between the lateral neural tube and the tip of the dermamyotome (Fig. 5.4). This region has been termed the migration “staging area” (Weston, 1991). Cells that enter this pathway will differentiate into pigment

Regulation of Melanoblast Migration and Differentiation +24 hrs

+24 hrs

Fig. 5.4. Schematic diagram of the major migratory pathways followed by neural crest cells in the trunk of amniote embryos. Anterior is to the left. (Bottom) Sagittal section through somites -6 to -12 indicating the simultaneous development of the myotome and neural crest invasion into the somite from the ventromedial path. (Top) Cross-sections at different axial levels show the ventromedial and dorsolateral pathways taken by neural crest cells. Diagram by Martha Spence and used with permission from Academic Press.

A

P

-12

-6

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cells of the skin. Melanoblasts begin entering the ectoderm approximately two stages after entering the dorsolateral path (Erickson et al., 1992). Eventually, they invade the feather follicle and complete their differentiation into melanocytes (Hulley et al., 1991). A major question about neural crest morphogenesis is what determines the positioning of neural and glial cells in the ventral pathway and pigment cells in the dorsolateral path. The generally accepted model for patterning neural crest derivatives proposes that neural crest cells are a pluripotent population of cells that migrate haphazardly into the ventral or dorsolateral paths, and differentiate according to the cues encountered in these paths (Le Douarin, 1982, 1986; Le Douarin et al., 1993; Selleck et al., 1993; Stemple and Anderson, 1993). This model is supported by experimental evidence from heterotopic grafting experiments (Le Douarin and Teillet, 1974; Le Douarin et al., 1975), as well as backtransplantation of crest-derived structures into the early migratory crest pathways (Ayer-Le Lievre and Le Douarin, 1982; Dupin, 1984; Le Douarin et al., 1978; Le Lievre et al., 1980), which show that, as a population, neural crest cells migrate and differentiate according to their new environment and not their origin. However, an alternative mechanism directs the migration of chicken melanoblasts into the dorsolateral path at the trunk level. Speciﬁcally, melanoblasts are developmentally biased at the time that they depart from the neural tube: only neural crest cells that are speciﬁed as melanocytes enter the dorsolateral path (Reedy et al., 1998a), and melanoblasts are the only neural crest cells that can exploit the dorsolateral path under experimental conditions (Erickson and Goins, 1995). Thus, the ﬁnal distribution of pigment cells is directed by cellautonomous migratory properties unique to that subpopulation. The evidence that melanoblasts are speciﬁed prior to entering the dorsolateral pathway will be discussed in the following section. Here we will ﬁrst consider what controls the choice of pathway. Why are neural and glial precursors unable to invade the dorsolateral path, whereas the later-migrating melanoblasts can exploit this pathway? The dorsolateral path at the trunk level is initially refractory to neural crest cell migration (Erickson and Goins, 1995; Erickson et al. 1992; Serbedzija et al., 1989). By stages 19–20, beginning at the wing-bud level, dorsolateral migration is initiated as ventral migration gradually ceases (Kitamura et al., 1992; Oakley et al., 1994; Serbedzija et al., 1989). The molecular basis for this switch in pathways, initially suggested by Oakley et al. (1994), was that inhibitory molecules are gradually lost, so that the environment in the dorsolateral path is now permissive for migration. However, experimental studies do not support this proposal. They instead suggest that there are changes in those neural crest cells speciﬁed as pigment cells that allow them to overcome the inhibitory cues. When nonmelanogenic neural crest cells are grafted into stage-19 embryos at the wing-bud level, which is the time when the host neural crest cells would be embarking on the dorsolateral path, the grafted cells only 120

migrate ventrally (Erickson and Goins, 1995). However, when melanoblasts are grafted into the early neural crest migratory pathway (stage-15 host), the grafted cells invade the dorsolateral space immediately, well in advance of the time when dorsolateral migration usually takes place (Erickson et al., 1992). These experimental studies indicate that molecular changes occurring during melanoblast speciﬁcation, rather than changes in the environment, permit dorsolateral migration. One possible molecular change in melanoblasts is that they develop unique sensitivity to positive guidance cues from the dermamyotome, allowing them to migrate dorsolaterally. In the mouse, melanoblasts depend upon SLF produced by the dermamyotome for their initial dispersal onto the dorsolateral path (Wehrle-Haller and Weston, 1995, 1997; Wehrle-Haller et al., 1996; and see Mammals below), and they express the receptor for SLF, kit, prior to embarking on the dorsolateral path. However in the avian embryo, kit is ﬁrst expressed by melanoblasts long after they have migrated dorsolaterally (stage 25), and SLF is not produced by the dermamyotome, but rather by the ectoderm and only after stage 25 (Lecoin et al., 1995; Reedy et al., 2003). Furthermore, morpholino knockdown of kit does not affect either melanoblast migration or speciﬁcation (Harris and Erickson, unpublished data). Thus, acquiring responsiveness to SLF owing to kit upregulation is unlikely to control the timing of dorsolateral migration in the chicken. In the past few years a number of guidance molecules have been identiﬁed that cause the collapse or repulsion of growth cones, ﬁlopodia or lamellipodia (Tessier-Lavigne and Goodman, 1996). These include the netrins, the collapsin/ semaphorins, Slits and ephrins (Gale and Yancopoulos, 1997). Three laboratories have shown that ventrally migrating neural crest cells are constrained to migrate in the anterior half of somites because inhibitory ephrins are expressed in the posterior somite (Flenniken et al., 1996; Krull et al., 1997; Wang and Anderson, 1997). Ephrins are also distributed in the dorsolateral space at the time of early crest migration and they have also been shown to be responsible for preventing neural and glial precursors from entering this space (Santiago and Erickson, 2002). In contrast, ephrins stimulate melanoblast migration, and this positive guidance cue permits melanoblasts to invade the dorsolateral space. This is not unlike the netrins and semaphorins, which can be inhibitory under some circumstances and stimulatory under others. A major task will be to understand how Eph receptors, the receptors for ephrins, can differentially signal in melanoblasts and nonmelanoblasts. Further, we do not yet understand how ephrin signaling results in enhanced melanoblast migration, although integrin activation is likely to be a factor (Beauvais et al., 1995; Santiago and Erickson, 2002). Ephrins are likely not the only signaling molecules that control the directional migration of melanoblasts into the dorsolateral space. Recently, Jia et al. (2005) have demonstrated that Slits, which are secreted chemorepellants, expressed in the dermamyotome also prevent the migration of early migratory neural crest cells into the dorsolateral space. When the Slit

REGULATION OF MELANOBLAST MIGRATION AND DIFFERENTIATION

receptor, Robo 1, is inactivated by expressing a dominant negative form in the premigratory neural crest, ventrally migrating crest cells now precociously invade the dorsolateral pathway. Still unanswered is how melanoblasts can overcome the chemorepellant action of Slit. Potentially the Robo receptors could be downregulated on neural crest cells during melanoblast speciﬁcation, or melanoblasts may be attracted to Slits rather than being repelled. Yet another signaling molecule/receptor pair that potentially controls directed migration into the dorsolateral space is endothelins/endothelin receptors (reviewed by Pla and Larue, 2003). Endothelins are 21-amino acid peptides originally identiﬁed for roles in vasoconstriction (Yanigasawa et al., 1988) and their receptors are G-protein-coupled, heptahelical, cellsurface receptors. In the chick, ventrally migrating neural crest cells express Ednrb, whereas Ednrb2 is upregulated just prior to neural crest cells migrating in the dorsolateral pathway. Endothelin-3, the presumptive ligand for avian Ednrb and Ednrb2, is expressed by the ectoderm (Nataf et al., 1998). When mouse embryonic stem (ES) cells are transfected with Ednrb2 and grafted into an early chick embryo, they take the dorsolateral pathway precociously, unlike the wild-type ES cells or ES cells expressing Ednrb, which migrate ventrally (Pla et al., 2005). This study suggests that endothelin signaling may positively attract melanoblasts into the dorsolateral space. It would be interesting to manipulate the composition of endothelin receptors on neural crest cells in situ to test this possibility directly. Beyond effects on migration, several studies of cultured quail neural crest cells show that endothelin-3 clearly affect proliferation of melanoblasts (Lahav et al., 1996). Additionally clonal analysis reveals that endothelin signaling enhances the survival of clonogenic cells that give rise to melanocytes only, glial cells only, and the melanocyte/glial bipotent precursors (Lahav et al., 1998). During later morphogenesis, melanoblasts enter the ectoderm, which is separated from the dermis by a basal lamina. Although previous studies have shown that basal laminae are impenetrable by early-migrating neural crest cells, melanoblasts clearly have enhanced invasive properties (Cuddenec, 1977; Erickson, 1987; Erickson and Goins, 1995; Sears and Ciment, 1988; Teillet, 1971). Localized discontinuities in the basal lamina adjacent to neural crest cells in the dorsolateral pathway suggest that melanoblasts selectively degrade the basal lamina prior to invading the ectoderm (Erickson et al., 1992). The mechanisms of melanoblast invasiveness are not clear, but one possibility is that they produce matrix-degrading proteases that allow them access to sites where other crest cells normally cannot enter (Duong and Erickson, 2004; Erickson and Isseroff, 1986; Valinsky and Le Douarin, 1985).

Differentiation of Avian Melanocytes The melanoblast lineage was believed to be speciﬁed only after neural crest cells had migrated from the neural tube, because a number of cloning or culture studies using avian neural crest showed that pigment cells could arise from a pluripotent pre-

cursor in 24-hour outgrowths (Barofﬁo et al., 1991; Dupin and Le Douarin, 2003; Le Douarin et al., 1993; Sieber-Blum et al., 1993; Stocker et al., 1991; Weston, 1991). However, when quail trunk-level neural tubes are cultured and the fate of those neural crest cells that emigrated during the ﬁrst six hours is assessed, no melanocytes differentiate, even after six days in culture under conditions that are permissive for melanogenesis (Reedy et al., 1998a). Thus there is heterogeneity in the melanogenic potential of different temporal subpopulations of neural crest cells from the same axial level. Corroborating studies by Henion and Weston (1997) showed that many of the neural crest cells are fate-restricted (that is, the progeny from a single neural crest cell all have the same phenotype) at the time that they detach from the neural tube, and that neurogenic and some gliogenic precursors migrate from the neural tube before any melanogenic precursors. They did note that there are some clones that are bipotent and could give rise to glial cells and pigment cells, which are likely to be the intermediate precursor population identiﬁed in previous cultures studies (Sherman et al., 1993; Sieber-Blum and Cohen, 1980). Together these two studies show that many neural crest cells are developmentally restricted when they leave the neural tube, and that neurogenic, gliogenic, and melanogenic neural crest cells are speciﬁed at different times prior to their migration. Similar observations have been made in the zebraﬁsh (Raible and Eisen, 1994; Dorsky et al., 1998, 2000a) and in axolotls (Epperlein and Löfberg, 1984). See also reviews by Anderson (2000), Dorsky et al. (2000b), and Erickson (2003). We know a great deal about melanogenesis, including the biochemical pathways that generate pigment, the regulation of expression of the enzymes in that pathway, and the role of numerous growth factors that play a role in melanoblast maintenance, proliferation, and terminal differentiation (discussed in detail in many chapters in this textbook). Nevertheless, very little is known about the molecular events that specify the lineage as distinct from the other neural crest derivatives. The studies cited above suggest that there is a lineage switch from the nonmelanogenic lineages to the melanogenic lineage; that is, the premigratory neural crest cells may be speciﬁed ﬁrst as neuro/gliogenic cells [or bipotent neural/glial and glial/melanogenic precursors (Le Douarin et al., 2004)] and subsequently some switch to the melanogenic fate. Further, this lineage switch occurs while the neural crest precursors are still resident in the neural tube or shortly after they detach from it. Studies in the chick (Jin et al., 2001) and in zebraﬁsh (Dorsky et al., 1998, and see Fishes, above) show that this lineage switch is controlled, at a minimum, by two competing classes of signaling molecules: the Wnts and the BMPs. Speciﬁcally, Wnt 1/3a signaling speciﬁes melanocytes whereas BMP 2/4 signaling speciﬁes neural and glial cells. Despite the evidence that Wnt and BMP signaling is involved in fate speciﬁcation, we do not know the cellular mechanisms by which this speciﬁcation is achieved, or how Wnt- and BMPdependent signals compete with each other in this context. There are undoubtedly many genes involved in the speciﬁcation of the melanogenic lineage downstream of Wnt 121

CHAPTER 5

signaling, including transcription factor cascades (Anderson, 1994; Groves and Anderson, 1996; Groves et al., 1995). Only a few transcription factors have been identiﬁed to date that regulate the melanogenic fate. Microphthalmia (Mitf), a basichelix-loop-helix-zipper transcription factor, has been shown in human, mouse, zebraﬁsh (nacre, mitfa), and chickens to bind to the so-called M-box, an 11-bp sequence containing the core element CATGTG (Hodgkinson et al., 1993), in the upstream regulatory region of the tyrosinase and tyrosine-related protein 1 (TRP-1) genes and activate their transcription (review by Goding, 2000; see Mammals, below, for more detail). Downregulation of Mitf in chick neural crest cells using morpholinos eliminates melanoblasts and misexpression of Mitf by electroporation gives rise to excess pigments cells (Thomas and Erickson, unpublished). Mitf appears to be the primary transcription factor that drives melanoblast speciﬁcation (Goding, 2000). Speciﬁcation of different neural crest subpopulations at the trunk level occurs in sequential temporal waves, and yet many transcription factors that control neural, glial, and melanogenic lineages are expressed at the same time in the dorsal neural tube. For example, Sox10 and Pax3 are both required for the development of all three lineages in mouse, chick, and probably zebraﬁsh (Britsch et al., 1998; Cheung et al., 2005; Dutton et al., 2001; McKeown et al., 2005; Lacosta et al., 2005; Potterf et al., 2001), so what prevents neuroblasts, glioblasts, and melanoblasts from developing simultaneously? One possibility is that a transcriptional repressor for melanogenesis is expressed early, which would prevent the early migratory crest from having melanogenic potential, whereas Wnt signaling may cause the downregulation of this putative repressor, which would then permit melanogenesis. We predict that the lineage switch from neural/glial to melanoblast occurs as a result of the downregulation of this repressor. A good candidate for such a transcriptional repressor is FoxD3. FoxD3 is a member of the winged-helix family of transcription factors (Kaestner et al., 2000). FoxD3 represses melanogenesis but is required for gliogenesis in the chick (Kos et al., 2001). The zebraﬁsh homolog (formerly, fkh-6) similarly stimulates glial cell differentiation (Kelsh et al., 2000) and foxd3 mutants exhibit defects in neurons, glia, and ectomesenchymal lineages, though not chromatophore lineages (Stewart et al., 2005). In the neural epithelium, FoxD3 controls another lineage switch from neural crest to interneuron (Dottori et al., 2001), and in the early mouse embryo it is critical for maintaining embryonic stem cells (Hanna et al., 2002). Since FoxD3 in most experimental contexts acts as a transcriptional repressor (Freyaldenhoven et al., 1997b; Hromas et al., 1999), the most parsimonious hypothesis that accounts for the current data is that FoxD3 represses Mitf expression directly by binding to the Mitf promoter. Using a luciferase assay, chick FoxD3 expressed in B16 melanoma cells will reduce luciferase activity that is being driven by a chicken Mitf enhancer (Thomas and Erickson, unpublished results). This model further predicts that when FoxD3 is downregulated under the inﬂuence of Wnt signaling, Mitf is no longer repressed 122

by FoxD3, and is therefore transcribed by the transcriptional activators Tcf/Lef-1, Sox10, and Pax3 (Goding, 2000).

Mutants There is considerable variation in plumage and skin pigmentation among domestic fowl (reviewed in Smyth, 1990; Staples, 2001) and a variety of mutants with melanocyte defects have been isolated (e.g., Niwa et al., 2002) including a mutation in Mitf (Mochii et al., 1998). Congenital hypopigmentation resulting from inefﬁcient melanoblast migration, differentiation, or both also has been shown in several instances (Jimbow et al., 1974). Some chicken strains have been used as model systems for studying pigment defects such as vitiligo (Boissy and Lamoreux, 1988). Most of these defects are characterized by the spontaneous depigmentation of normally pigmented birds and probably represent mutants in melanocyte physiology rather than true developmental mutants. One exception is the Silkie strain of chickens, which is characterized by extensive dermal and internal pigmentation caused by a prevalence of melanocytes in ectopic locations (Faraco et al., 2001; Hallet and Ferrand, 1984; Makita and Mochizuki, 1984; Makita and Tsuzuki, 1986). Unlike White Leghorn and quail melanoblasts, Silkie melanoblasts migrate ventrally, as well as dorsolaterally, during neural crest morphogenesis (Faraco et al., 2001; Reedy et al., 1998b). The abnormal migration of melanocytes in Silkie chickens does not reﬂect a defect in the neural crest, because Silkie neural tubes grafted into White Leghorn hosts never exhibit internal pigmentation, whereas quail neural tubes placed in Silkie embryos do generate internal melanocytes (Hallet and Ferrand, 1984). A molecular marker for barrier molecules, PNA (Oakley et al., 1991), indicates that inhibitory molecules are normally distributed beneath the dermamyotome in regions through which melanoblasts do not migrate. These PNA barriers are absent in the Silkie embryo (Faraco et al., 2001; de Freitas et al., 2003), suggesting that barriers develop during the late migration of melanoblasts that keep them in the dorsolateral path and prevent them from dispersing ventrally. Thus this mutant has identiﬁed a process late in melanoblast migration that constrains melanoblasts to the skin. Identiﬁcation of the factors expressed in the Silkie environment responsible for melanocyte morphogenesis will be of considerable interest.

Mammals As in birds, the only pigment cell found in mammalian skin is the melanocyte. The neural crest origin of mammalian epidermal melanocytes was demonstrated by Rawles (1947, 1948) in an elegant study in which she grafted isolated mouse neural tube, somite, or skin (ectoderm plus dermis) of varying ages into the chick coelom and then scored for the presence of pigmented cells. Studies of mammalian development are hampered by several factors, including the inaccessibility of the embryo to experimental manipulation and higher maintenance costs compared with other vertebrate systems. Nevertheless, the mouse has proved to be a popular system for

REGULATION OF MELANOBLAST MIGRATION AND DIFFERENTIATION

studying pigment cell development because of the ability to make and analyze transgenic embryos and the availability of many coat color mutants. In this section we discuss the development of pigmentation in mammals, focusing primarily on mouse and to a lesser extent on humans.

Morphogenesis of Murine Melanocytes Several studies have followed the early stages of trunk neural crest migration in mouse embryos (e.g., Erickson and Weston, 1983; Serbedzija et al., 1990). At the level of the forelimb, neural crest cells ﬁrst emerge from the neural tube at day E8.5 (i.e., 8.5 days postcoitus). As in avian embryos, mammalian trunk neural crest cells take one of two migratory pathways: ventrally between the somite and neural tube, or dorsolaterally in the space between the dermamyotome and overlying somite (Rawles, 1947; Serbedzija et al., 1990). Unlike the situation in avians, however, the entry of mouse neural crest cells into the dorsolateral path does not appear to be delayed signiﬁcantly with respect to ventral migration (Rawles, 1947; Serbedzija et al., 1990, 1994). By E10.5, the emigration of neural crest cells from the dorsal neural tube has ceased (Serbedzija et al., 1990, 1994). Mayer (1973) recombined ﬂank dermis and ectoderm from albino or normally pigmented mouse embryos to demonstrate that melanoblasts begin entering the epidermis at E12. This was subsequently conﬁrmed using probes against the melanogenic enzyme dct to identify melanoblasts (Cable et al., 1995; Pavan and Tilghman, 1994; Steel et al., 1992). Interestingly, the ﬁrst melanoblasts to enter the ectoderm are located lateral, as opposed to dorsal, to the neural tube (Pavan and Tilghman, 1994). This corresponds to the spatial pattern of the onset of dct immunoreactivity and suggests that entry into the epidermis depends on the cells reaching a particular point in melanogenesis. Once in the epidermis, melanoblasts migrate into the developing hair follicles, and by E16 premelanosomes are detectable by the dopa reaction (reviewed in Hirobe, 1995). More recently, several studies have used mice that express the lacZ reporter under the control of the Dct promoter/ enhancer elements to track murine melanoblasts more carefully. For example, Wilkie and coworkers (2002) used Dct-lacZ transgenic mice to study the spatial and temporal patterns of melanoblast appearance and migration. They noted that melanoblasts do not arise uniformly along the length of the neural tube. Rather, a large pool of melanoblasts is present in the cervical region by E11.5, and signiﬁcantly fewer melanoblasts are found in the trunk. The cervical pool appears to be the source of craniofacial melanocytes, as no melanoblasts were found to originate in rostral regions. Also using Dct-lacZ reporter mice, Jordan and Jackson (2000b) found evidence that there is a kit-independent population of melanoblasts present in the trunk between E15.5 and E16.5, and that these cells are capable of colonizing areas of the ﬂank that are otherwise devoid of melanoblasts in the Ph mutants. The ontogeny of these cells is as yet unclear, but if conﬁrmed this is a very interesting observation because of the

obvious parallels reported in zebraﬁsh (discussed in Fish, above). Distribution and patterning of pigment cells is controlled not only by guidance molecules but also by signaling systems that control cell proliferation and cell survival. In the mouse there is, as yet, little information about the molecular nature of directed migration of melanoblasts. However, two general classes of mouse mutants have identiﬁed critical regulators of melanoblast proliferation and survival that in turn affect patterning: piebald/lethal spotting and Steel/Dominant spotting. piebald mutants display a range of white spotting depending on the allele. Only neural crest-derived melanocytes are affected, however, as retinal pigmentation is normal (Silvers, 1979). In addition to white spotting, piebald mutants exhibit dramatically reduced numbers of two other neural crest derivatives: the cochlear melanocytes of the inner ear, and the enteric ganglia of the myenteric plexus (Deol, 1967, 1970; Lane, 1966; Silvers, 1979; Webster, 1973). In homozygotes of the most severe allele, piebald-lethal (sl), this results in deformities of the cochlear duct and in megacolon, the latter of which causes neonatal death. The product of the s locus was identiﬁed as the endothelinB receptor (Ednrb) (Greenstein et al., 1994; Hosoda et al., 1994). Analysis of the Ednrb gene in piebald mutants provides an explanation for the variation in allelic severity; whereas the entire Ednrb gene is deleted in the sl allele, the less severe s allele is structurally normal but expressed at much lower levels compared to wild type (Hosoda et al., 1994). Interestingly, early tissue recombination experiments (Mayer 1965, 1967a, b, 1977) suggested that although the s phenotype is largely cell autonomous, there were non-cell autonomous effects as well. More recent tissue recombination experiments by Lou and coworkers (2004) support this conclusion. Pavan and Tilghman (1994) demonstrated that the white spotting phenotype in sl mutants is manifested very early in development by an almost complete lack of melanoblasts in the trunk. Experiments by Shin and coworkers (1999) using a conditional allele conﬁrmed that melanocytes are dependent upon Ednrb very early in their development, between E10 and E12, consistent with Ednrb expression in the head fold and future dorsal neuroepithelium prior to neural crest dispersal (Tilghman et al., 1995). What happens to melanocyte precursors in the absence of Ednrb function? Recent studies by Lee et al. (2003) have shown that melanoblasts form and even begin to disperse from the neural tube in Ednrb-null embryos. However, Ednrb-null melanoblasts do not migrate very far along the dorsolateral pathway and eventually disappear. Whether these cells ultimately die or simply fail to complete melanogenesis remains unclear. It should also be noted that in the chicken embryo (see above), melanoblasts chemotact toward endothelin-3. Whether endothelins provide directional cues to murine melanoblasts has not been explored. The identiﬁcation of Ednrb as the gene at the piebald locus suggested that one or more of the endothelins might play a role 123

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in melanocyte development. The phenotype of mutants at the lethal spotting (ls) locus is virtually identical to that of piebald mutants, including white spotting and megacolon (Silvers, 1979). Baynash et al. (1994) demonstrated that the product of the ls locus is EDN-3. In addition, they showed that ls mutants carry a point mutation that prevents normal proteolytic processing of the inactive polypeptide precursor of EDN-3. The other mouse mutants that have provided important insights into morphogenesis are Dominant spotting (W) and Steel (Sl). W and Sl mutants are phenocopies of one another and are characterized by being sterile, anemic, lacking in mature mast cells, and devoid of epidermal pigmentation (Silvers, 1979). These deﬁciencies are caused by the abnormal development of three stem cell populations: the germ cells, hematopoietic precursor cells, and the neural crest [Silvers (1979) and references therein; reviewed in Fleischman (1993), Morrison-Graham and Takahashi (1993)]. Early work demonstrated that the Sl mutation, but not the W mutation, could be rescued by a “normal” environment, implying that the W mutation was cell autonomous whereas the Sl mutation was not (Mayer, 1970; Mayer and Green, 1968). These results, together with the complementary mutant phenotypes, suggested that the W and Sl gene products act in a common pathway, possibly as a receptor–ligand complex. The cloning and characterization of these genes proved that this is, in fact, the case, with W encoding Kit and Sl encoding the Kit ligand, SLF (Anderson et al., 1990; Flanagan and Leder, 1990; Geissler et al., 1988; Huang et al., 1990; Martin et al., 1990; Williams et al., 1990; Zsebo et al., 1990a, b). Several studies have attempted to deﬁne exactly when melanoblasts require Kit and SLF activity (see Wehrle-Haller, 2003 for a comprehensive review). The patterns of expression of both Kit and SLF mRNAs suggest an early role in melanocyte development, with Kit+ melanoblasts detectable in the trunk as early as E10.5 (Keshet et al., 1991; Manova and Bacharova, 1991; Wehrle-Haller and Weston, 1995). Likewise, in utero injection of ACK2, a Kit function-blocking antibody, revealed that Kit is required at this time (Nishikawa et al., 1991; Yoshida et al., 1993). These results are consistent with several studies that examined the dependence of melanogenesis on SLF in vitro (Morrison-Graham and Weston, 1993; Murphy et al., 1992; Reid et al., 1995). In vitro and in vivo studies support the idea that mammalian Kit signaling is required for the survival of melanoblasts and their proper migration (Cable et al., 1995; Ito et al., 1999; Morrison-Graham and Weston, 1993; Murphy et al., 1992; Reid et al., 1995; Steel et al., 1992; Wehrle-Haller and Weston, 1995; Wehrle-Haller et al., 2001; reviewed in Yoshida et al., 2001) as in ﬁsh (Parichy et al., 1999; Rawls and Johnson, 2003). Later in development, Kit/SLF signaling is also required for the entry of melanoblasts into the developing hair follicle, having largely chemokinetic effects on melanoblasts (Jordan and Jackson, 2000a; Peters et al., 2002); the precise factors that guide these precursors to hair follicles remain unknown. Remaining unresolved is the importance of two forms of 124

SLF. Alternative splicing of the primary Sl transcript results in two isoforms, soluble and membrane bound, both of which are equally capable of binding to and activating Kit (Flanagan et al., 1991; Majumdar et al., 1994). In Sl dickey (Sld) mutants, the entire transmembrane and cytoplasmic domains of the gene are deleted. As a result, Sld mutants produce functional, soluble SLF but no transmembrane SLF (Brannan et al., 1991; Flanagan et al., 1991). Sld/Sld homozygotes are phenotypically identical to homozygotes for Sl null alleles (Silvers, 1979), indicating that soluble SLF alone is not sufﬁcient for normal melanogenesis. Wehrle-Haller and Weston (1995) demonstrated that soluble SLF is, however, sufﬁcient for the initial dispersal of melanoblasts along the dorsolateral pathway, while membrane-bound SLF is required for later survival, differentiation, or both. One explanation for the importance of membrane-bound SLF is that it would be expected to result in a higher local concentration of SLF because it cannot diffuse away from the cells producing it. An additional possibility, however, is that the membrane-bound form of SLF functions as a cell adhesion partner for Kit expressing melanoblasts. The ability of SLF/Kit binding to promote cell adhesion directly has been demonstrated in cell lines (Flanagan et al., 1991), primordial germ cells (Marziali et al., 1993), mast cells (Adachi et al., 1992; Kaneko et al., 1991), and megakaryocytes (Avraham et al., 1992).

Morphogenesis of Human Melanocytes Despite the evidence for a neural crest origin for pigment cells in other vertebrate species, many early studies of human pigmentation were prefaced on the theory that human melanocytes are derived from the basal cells of the epidermis, rather than the neural crest [Bloch (1917, 1921), as reported in Zimmerman and Becker (1959a)]. Information on human melanocyte development is reviewed in Holbrook et al. (1988) and Boissy and Nordlund (1995) and is summarized brieﬂy here. The migration of neural crest through the lateral pathway brings melanoblasts ﬁrst to the dermis. From the dermis, melanoblasts migrate into the epidermis to their eventual location on the basal lamina of the basal epithelial layer of the epidermis. Early work on the timing of appearance of fetal melanoblasts and melanocytes was limited to indirect approaches that utilized the reactivity of premelanosomes with reduced silver and subsequent identiﬁcation of reactive cells by electron microscopy. Mature melanocytes have also been identiﬁed using the dopa reaction and morphologic criteria. More recently, the discovery of a cytoplasmic epitope shared by melanoma cells and fetal melanocytes, the HMB-45 monoclonal antibody (Gown et al., 1986), has allowed a reanalysis of data on the timing of appearance of human melanocytes (Holbrook et al., 1988). Early studies indicated that in human embryos neural crest migration is complete by seven weeks and that melanoblasts appear in the dermis by 11 weeks (Zimmerman and Becker, 1959a, b). The timing of appearance of melanoblasts in the epidermis has since been reﬁned to earlier than seven weeks using HMB-45 staining (Holbrook et al., 1988). The density

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of this initial population of melanocytes observed by seven weeks is about 50% of the density observed at birth. By 10 weeks, melanocytes in the epidermis are observed to have premelanosomes based on both the reaction with reduced silver and direct electron microscopic observation (Fujita et al., 1970; Zimmerman and Becker, 1959b). It was thought that a large inﬂux of melanoblasts occurred at the 12–14-week stage of development based on counts derived by reduced silver staining, but this has been reinterpreted by the HMB-45 staining data, which seem to imply that the levels of melanocytes thought to appear at the twelfth week are already in place by the eighth week and thus there is no new inﬂux. There is a rough doubling of the number of melanocytes between weeks 10 and 14, with the numbers observed by HMB-45 staining much greater than the numbers observed by reduced silver staining. This may be due to mitosis of melanocytes that are already in the epidermis. The ﬁnal number and distribution of melanocytes in the adult is relatively constant, although it can be affected by such stimuli as ultraviolet (UV) light. The distribution of melanocytes appears to be nonrandom (Holbrook et al., 1988), and each melanocyte is found with 36 keratinocytes in an epidermal melanin unit. Melanocytes that are associated with hair follicles arrive at their ﬁnal location by following the downgrowth of epidermal cells in the developing hair germ (Mishima and Widlan, 1966). The production of melanin pigment developmentally begins at around 12 weeks based on both electron microscopic studies and dopa staining (Zimmerman and Becker, 1959a; Zimmerman and Cornbleet, 1948). The transfer of pigment to keratinocytes occurs late in the second trimester of development (Zimmerman and Cornbleet, 1948). The human piebald trait, characterized by a white forelock and patchy pigmentation, is likely due to a defect in melanocyte morphogenesis resulting in reduction or absence of these cells in discrete areas of the skin. Like white spotting in mice, piebaldism is congenital (Halaban and Moellmann, 1993) and many piebald patients have been shown to have mutations in the human homolog of Kit (e.g., Fleischman, 1992, 1993; Fleischman et al., 1991; Giebel and Spritz, 1991; Spritz et al., 1992, 1993; reviewed in Baxter et al., 2004). Defects in melanocyte and melanoblast morphogenesis resulting from disruptions in endothelin-3 and Ednrl signaling are likewise implicated in Hirschsprung disease in humans, which is also characterized by aganglionic megacolon (McCallion and Chakravarti, 2001).

Differentiation of Mammalian Melanocytes As discussed previously, it is well documented in both teleost and avian embryos that melanocyte lineage segregation begins very early in neural crest development, and that melanoblasts are speciﬁed prior to or at least coincident with their emigration from the neural tube. A recent study by Wilson et al. (2004) demonstrates that the same is true in murine embryos. Using mice that express lacZ under the control of the kit regulatory elements (Wlacz), they identiﬁed distinct melanogenic (kit+) and neurogenic (p75+) subpopulations of

premigratory neural crest cells in the midline of the dorsal neural tube prior to neural crest emigration. What signaling pathways are responsible for the initial segregation of the melanocyte lineage? There is strong evidence to suggest that Wnt/b-catenin signaling plays a critical role in this process. Wnt signaling promotes melanocyte formation in both birds and ﬁsh (see previous sections), and targeted disruption of the b-catenin gene in the dorsal neural tube of mouse embryos results in speciﬁc loss of the melanocyte and sensory ganglia neural crest subpopulations (Hari et al., 2002). The most likely activators of b-catenin in vivo are Wnt1 and Wnt3a, both of which are expressed in the dorsal neural tube and required for melanocyte formation (Ikeya et al., 1997). Dunn et al. (2005) have recently shown that forced overexpression of b-catenin, Wnt1, or Wnt3a in vitro results in an endothelin-dependent increase in melanocyte differentiation as well. Several transcription factors have been identiﬁed that are required for neural crest development overall (e.g., slug/snail, foxd3, and Pax 3) and are therefore important for melanocyte formation as well. The ﬁrst transcription factor that is relatively speciﬁc for melanocyte differentiation was identiﬁed in the microphthalmia (mi) mutant. Both neural crest-derived and non-neural crest-derived melanocytes are affected in mi mutants. As a result, homozygotes of many alleles display white spotting, small, unpigmented eyes, and a lack of melanocytes in the inner ear (Silvers, 1979). Mice with certain mi alleles are also deﬁcient in osteoclasts and mast cells (Silvers, 1979). mi encodes the transcription factor Mitf (Hodgkinson et al., 1993). Members of this family bind DNA as homo- or heterodimers (Hemesath et al., 1994), which explains why the original mi allele is semidominant and may also explain the high degree of intrallelic complementation and interallelic interactions that characterize mi mutants (Hodgkinson et al., 1993; Silvers, 1979; Steingrimsson et al., 1994). As previously discussed, Mitf regulates the expression of several melanocytespeciﬁc genes, including dopachrome tautomerase, tyrosinase, tyrosinase-related protein 1, Pmel17, and Melan-a (Baxter and Pavan, 2003; Bentley et al., 1994; Du et al., 2003; Yasumoto et al., 1995; Yokoyama et al., 1994). In transgenic mice carrying mutations in mi, melanoblasts form initially but fail to survive and reach the migration staging area (Hornyak et al., 2001). It is unlikely that this phenotype is related to Mitf’s role regulating expression of the melanin biosynthesis genes mentioned above, as mutations at the brown (TRP-1), slaty (dct), and albino (tyrosinase) loci do not result in melanocyte death but rather prevent the formation of normal melanin (see Silvers, 1979; Bennett, 1993; and references therein). However, McGill et al. (2002) have shown that Mitf is a critical transcriptional regulator of the antiapoptotic gene Bcl2 in melanocytes, suggesting that Mideﬁcient melanoblasts die as a result of Bcl2 downregulation. Mitf can activate transcription in vitro through the M box found in the promoters of several melanocyte-speciﬁc genes (Bentley et al., 1994; Lowings et al., 1992; Shibahara et al., 1991; Yasumoto et al., 1994; Yavuzer and Goding, 1994). 125

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Moreover, misexpression of Mitf can convert nonmelanocytes into melanin-producing cells and rescue melanogenesis in mutant mice (Opdecamp et al., 1997). As in other vertebrates, Mitf appears to be one of the major links between Wnt signaling and melanogenesis in two ways. First, MITF is a direct target of Wnt3a-mediated LEF-1 activation (Takeda et al., 2000; see also Fishes). Second, the Mitf protein interacts in trans with LEF-1 to regulate expression of melanocyte-speciﬁc genes (Yasumoto et al., 2002). In humans, mutations in Mi are associated with Waardenburg’s syndrome type 2A (WS2A), an autosomal dominant condition exhibiting deafness and patchy, abnormal pigmentation (reviewed in Baxter et al., 2004; Hughes et al., 1994; Tassabehji et al., 1994b). The importance of Mitf in WS may be even more far reaching because two other genes linked to WS discussed below, sox10 and Pax3, regulate Mitf expression (Lee et al., 2000; Potterf et al., 2000; Verastegui et al., 2000). Thus, it is possible that the common etiologies between the various forms of WS ultimately result from a lack of Mitf expression. Sox10 is another transcription factor that is critical for melanocyte development (reviewed in Mollaaghababa and Pavan, 2003). Sox10 directly regulates the expression of MITF (Watanabe et al., 2002), and the sox10 protein also cooperates with the Mitf protein to regulate expression of Dct (Potterf et al., 2001; Ludwig et al., 2004). Sox10 is also associated with WS2A in humans. A third transcription factor in the melanogenesis regulatory network was identiﬁed in the Splotch mutant. The pigment defect in Splotch heterozygotes is characterized by white spotting on the feet, tail, and belly (Silvers, 1979). For most alleles, the homozygous phenotype is lethal due to defects in the neural crest and to the failure of the neural tube to close properly (Jackson, 1994; Silvers, 1979 and references therein). Other neural crest derivatives affected in homozygous splotch mutants include the spinal root ganglia and Schwann cells (Silvers, 1979). Interestingly, these derivatives are not noticeably affected in heterozygotes. The splotch locus corresponds to Pax3, encoding a member of the paired-box transcription factor family (reviewed in Chalepakis et al., 1993) that is expressed in the dorsal neural tube and early neural crest cells (Goulding et al., 1991). Melanoblasts do develop in splotch mice, but they are present in far lower numbers than normal (Hornyak et al., 2001). This may be due at least in part to the role of Pax3 in regulating expression of Mitf. Some patients with WS1 have been shown to have mutations in the human homolog of Pax3 (e.g., Tassabehji et al., 1992, 1994a). A recent report shows that Pax3 activates Mitf, while simultaneously competing with Mitf for sites in the dct promoter. Wnt signaling and b-catenin activation relieve this repression, allowing melanogenesis to proceed. Thus, Pax3 appears to prime cells for the differentiation whereas Wnt signaling allows the cells to proceed down this pathway (Lang et al., 2005).

Mutants Much of what we know about melanocyte development in mammals has come from studies of the over 70 mouse pigment 126

mutants. A detailed discussion of the development and initial characterization of these lines can be found in Silver’s classical 1979 treatise, The Coat Colors of Mice. In general, mouse coat color mutants can be assigned to one of three categories: those that affect the subcellular structure of melanocytes, those that perturb the biochemical pathways of melanin synthesis, and those that disrupt normal melanocyte development. Mutants in the latter category typically display various degrees of white spotting, i.e., an absence of melanocytes from distinct regions of the head, trunk, or rump. Comprehensive discussions of these coat color mutants can be found elsewhere (e.g., Baxter et al., 2004; Bennett, 1993; Jackson, 1994; Silvers, 1979). While many have been identiﬁed at the molecular level, much work remains to be done. For example, the patchwork mouse, which exhibits a salt-andpepper phenotype, may be especially interesting for understanding melanoblast colonization and survival in the hair follicle (Aubin-Houzelstein and Panthier, 1999; AubinHouzelstein et al., 1998). A ﬁnal mutant worth mentioning for historical reasons is Patch, a homozygous-lethal dominant mutation. Heterozygotes have a sharply demarcated white belly spot that is variable in size, sometimes encircling the entire trunk (Silvers, 1979). The Patch mutation was mapped to a deletion that encompasses the gene for platelet-derived growth factor receptor-a (Pdgfra) (Stephenson et al., 1991). The Pdgfra gene is linked to Kit on chromosome 5 (Nagle et al., 1994). Interestingly, the Patch phenotype (MorrisonGraham et al., 1992) is similar to that of KitW–sh mutants. The KitW–sh mutation is a gross rearrangement of the 5¢ upstream region of the Kit gene that results in the ectopic misexpression of Kit in a variety of tissues, including the dermatome and developing limb (Duttlinger et al., 1993). It is hypothesized that this misexpression results in a reduction of the amount of SLF available to migrating melanoblasts (Besmer et al., 1993; Duttlinger et al., 1993). The phenotypic similarity of KitW–sh and Patch, together with the fact that the two genes are closely linked, led to the hypothesis that the Patch phenotype is due to the misexpression of Kit, rather than a reﬂection of melanoblast or neural crest dependence on Pdgfra activity in vivo. This hypothesis is supported by the observation that Kit mRNA is expressed ectopically in the dorsal neural tube and dermatome of Patch heterozygotes, despite the fact that Kit is not affected structurally by the Patch deletion (Duttlinger et al., 1995). The absence of pigment defects in targeted Pdgrfa knockout mice is consistent with a primary role for Kit misexpression in explaining the Patch phenotype (Soriano, 1997).

Perspectives The study of pigment cell development has been an area of active research in developmental biology for over a century because pigment cells are so readily identiﬁable and pigment patterns vary strikingly both within and among vertebrate

REGULATION OF MELANOBLAST MIGRATION AND DIFFERENTIATION

taxa. Until recently, embryologists studying pigment cell development were limited to careful descriptive analyses or physical perturbations of developing embryos. Now, these techniques are complemented by a growing understanding of the factors at the molecular and cellular level that regulate the morphogenetic behavior and differentiation of pigment cells. A comparative approach to studying pigment cell development has been, and will continue to be, absolutely critical for a thorough understanding of the subject. Although each system has it own unique advantages, no single system is ideal for studying both the molecular and cellular aspects of neural crest and pigment cell development. For example, identiﬁcation of the molecular basis for some of the mouse coat color mutants has provided deﬁnitive evidence for speciﬁc factors involved in melanocyte development. Nevertheless, murine embryos typically are not an experimentally tractable system, so characterizing the cellular effects of these factors is difﬁcult. Conversely, amphibian and avian embryos are highly accessible to experimental manipulation, but are only beginning to emerge as useful systems for genetic or molecular analyses. For this reason zebraﬁsh has become a powerful model system to study pigment patterns since they are both genetically and experimentally amenable. Comparative studies of pigment cell development also are useful for studying questions of developmental biology in an evolutionary context. Pigment cells in all vertebrates are derived from the same progenitor cell population, the neural crest, and at least one type of pigment cell, the melanophore/cyte, utilizes a common pigment biosynthesis pathway across species. These commonalities, and considerable research over the last decade, suggest that speciﬁc molecules will play similar roles in the morphogenesis or differentiation of pigment cells in different taxa. Equally interesting, however, are those instances where this is not true. For example, the importance of Kit signaling in the early migration and survival of zebraﬁsh melanophores and murine melanocytes is well established. However, the expression patterns of kit in amphibians and birds suggest this gene is not involved in the same events as in other vertebrates (Baker et al., 1995; Lecoin et al., 1995; Reedy et al., 2003). Discrete cells (pigment cells) from a well-deﬁned progenitor cell population (the neural crest) will thus continue to provide intriguing experimental materials for cell and developmental biologists in their quest to understand how eukaryotic cells differentiate and evolve.

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Murdock, F. W. Jacobsen, K. E. Langley, K. A. Smith, T. Takeishi, B. M. Cattanach, S. J. Galli, and S. V. Suggs. Stem cell factor is encoded at the Sl locus of the mouse and is the ligand for the c-kit receptor. Cell 63:213–214, 1990a. Zsebo, K. M., J. Wypych, and I. K. McNiece. Identiﬁcation, puriﬁcation, and biological characterization of hematopoietic stem cell factor from buffalo rat liver-conditioned medium. Cell 63:195–201, 1990b.

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The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

6

Melanoblast Development and Associated Disorders Richard A. Spritz

As discussed in Chapter 29, genetic disorders of melanocyte development, such as piebaldism and Waardenburg syndrome, are associated with heterogeneous distribution of pigmentation resulting from abnormal distribution of melanocytes during embryogenesis. These disorders result from mutations in a complex network of interacting genes that regulate differentiation and development of the neural crest, and the melanoblast/melanocyte lineage in particular, in both man and the laboratory mouse. Indeed, elucidation of the molecular basis of these disorders has provided a paradigm for the use of human and mouse genetics in combination to dissect the molecular etiology of genetic diseases. In the following discussion of these human genetic abnormalities of melanocyte development, MIM numbers for each disorder are from Online Mendelian Inheritance in Man (2004).

Piebaldism (MIM #172800) As discussed in Chapter 29, piebaldism is an autosomal dominant disorder characterized clinically by congenital patches of white skin and white hair, principally located on the frontal scalp, forehead, ventral chest and abdomen, and extremities (cf. Campbell and Swift, 1962; Comings and Odland, 1966; Cooke, 1952; Froggat, 1951; Geerts-Marchandise and Lachapelle, 1977; Grupper et al., 1970; Jahr and McIntire, 1954; Keeler, 1934; Küster, 1987; Pearson, 1913; Smith and Schulz, 1955; Storkan and Hosmer, 1954; Sundfor, 1939; Taylor and Robinson, 1976; Winship et al., 1991), which typically lack most or all melanocytes (Breathnach et al., 1965; Comings and Odland, 1966; Fukai et al., 1989; Jimbow et al., 1975). Piebaldism is now known to result from mutations of the KIT gene, which encodes the cell surface receptor for a growth factor ligand that plays a key role in melanogenesis, gametogenesis, and the early stages of hematopoiesis. The critical clues to the molecular basis of human piebaldism came from studies of what ultimately proved to be the homologous disorder of mice (reviewed by Spritz, 1992, 1993; Spritz and Hearing, 1994). W mutant mice, with so-called dominant white spotting, exhibit patches of white skin and fur very similar to the patches of leukoderma and poliosis of human piebaldism, as well as the features of hypoplastic anemia and sterility (reviewed by Bennett and Lamoreux, 2003; Lyon and Searle, 1989; Silvers, 1979). Molecular genetic studies demonstrated that mouse dominant white spot140

ting results from deletions or point mutations of the c-Kit (ofﬁcially, Kit) protooncogene (Chabot et al., 1988; Geissler et al., 1988; Hayashi et al., 1991; Larue et al., 1992; Nocka et al., 1990; Reith et al., 1990; Tan et al., 1990; reviewed by Morrison-Graham and Takahashi, 1993). The Kit gene, originally identiﬁed in the genome of the HZ4-feline sarcoma virus (Besmer et al., 1986), encodes the cell surface receptor for an embryonic growth factor, usually called “stem cell factor” (SCF), although also variously referred to as “Steel factor” (SLF), “mast cell growth factor” (MCGF), and “Kit ligand” (KL) (Copeland et al., 1990; Flanagan and Leder, 1990; Huang et al., 1990; Murphy et al., 1992; Zsebo et al., 1990; reviewed by Morrison-Graham and Takahashi, 1993). SCF is the product of the Steel (Sl) locus of mice, and both Sl and W mutant mice display similar piebald-like phenotypes. The KIT receptor is expressed on the surface of melanocytes and, whereas Sl mutant melanocytes develop normally when transplanted to normal skin, because normal ligand is available, W mutant melanocytes do not, as they have an intrinsic defect of receptor function. The KIT protein is a member of the tyrosine kinase family of transmembrane receptors. As illustrated in Fig. 6.1, the KIT polypeptide consists of an amino-terminal, extracellular, ligand-binding, receptor domain composed of ﬁve immunoglobulin-type domain repeats, a short transmembrane domain, and an intracellular domain consisting of a bipartite tyrosine kinase domain followed by a carboxyl-terminal tail. Within the extracellular domain, the ﬁrst three immunoglobulin-type repeats bind the SCF ligand (Blechman et al., 1993; Jiang et al., 2000; Lev et al., 1993), the fourth may (or may not) be involved in receptor dimerization (Blechman et al., 1995; Lemmon et al., 1997), and the ﬁfth is required for proteolytic release of the KIT receptor from the cell surface (Broudy et al., 2001). The stoichiometry of interaction between SCF and the extracellular domain of the KIT receptor is apparently monovalent (Lev et al., 1992), each subunit of the SCF dimer binding to one KIT chain on the cell surface (Jiang et al., 2000; Lemmon et al., 1997; Philo et al., 1996) and thereby inducing dimerization of the KIT receptor within the cell membrane and consequent activation of its intracellular tyrosine kinase (Lev et al., 1992). This, in turn, results in autophosphorylation of speciﬁc tyrosine residues within the KIT kinase domain, enhancing the binding of various proteins, including phosphatidylinositol 3¢ kinase and phospholipase Cg1, which act as downstream mediators of the mitogenic signal in the KIT-dependent pathway of signal transduction

MELANOBLAST DEVELOPMENT AND ASSOCIATED DISORDERS

Fig. 6.1. Schematic diagram of the human KIT polypeptide and locations of mutations that are associated with human diseases. The amino-terminal extracellular leader sequence (LS) and penta-repetitive ligandbinding domain, the transmembrane domain (TM), and the intracellular bipartite tyrosine kinase domain are indicated. Missense amino acid substitutions are indicated by ﬁlled circles, frameshifts by ﬁlled diamonds, nonsense mutations by ﬁlled triangles, splice junction mutations by zig-zags, and in-frame deletions by brackets. AML, acute myelogenous leukemia; GIST, gastrointestinal stromal tumor.

(reviewed by Morrison-Graham and Takahashi, 1993), Src family members, the JAK/STAT pathway, and the Ras–Raf–MAP kinase cascade (reviewed by Ronnstrand, 2004; Taylor and Metcalfe, 2000). The speciﬁc mechanism by which reduced KIT function results in the piebald phenotype is not known exactly. Injection of anti-kit antibodies into developing mouse embryos resulted in W-like pigmentary anomalies (Nishikawa et al., 1991), and incubation of cultured human melanocytes with antisense KIT oligonucleotide signiﬁcantly reduced melanocyte proliferation but not cell survival (Spritz et al., 1994a). It is clear that, in the mouse and zebraﬁsh, and presumably also the human, KIT function is required for normal proliferation and survival of both melanoblasts and melanocytes both before and after migration from the neural crest (Mackenzie et al., 1997; Nishikawa et al., 1991; Parichy et al., 1999; Rawls and Johnson, 2003; Spritz et al., 1994a; reviewed by Yoshida et al., 2001). Reduced KIT-dependent signal transduction results in a greatly reduced number of dermal melanocytes in the regions of involved skin. Nevertheless, the precise basis for the aberrant regional distribution of skin melanocytes, and particularly the typical pattern seen in human piebaldism, remains unknown. The ﬁrst clue to the molecular genetic basis of human piebaldism came from the observation of several patients with sporadic, nonfamilial piebaldism associated with de novo chromosomal deletions involving region 4q12 (Funderburk and Crandall, 1974; Hoo et al., 1986; Lacassie et al., 1977; Yamamoto et al., 1989). The human KIT gene was subsequently mapped to approximately the same chromosomal location (d’Auriol et al., 1988; Yarden et al., 1987), and this

coincidence suggested that human piebaldism might, like mouse “dominant white spotting,” also result from abnormalities of the KIT gene. In fact, the human KIT gene consists of 21 exons spanning almost 83 kb at chromosome segment 4q12 (Giebel et al., 1992; Vandenbark et al., 1992) and is part of a gene cluster encoding a family of closely related “type III” receptor tyrosine kinases, organized PDGFRAKIT-KDR (Spritz et al., 1994b). PDGFRA encodes the platelet-derived growth factor receptor a chain, and KDR the so-called “vascular endothelial growth factor receptor 2” (VEGFR2). The ﬁrst direct evidence for the involvement of KIT mutations in human piebaldism was provided by Giebel and Spritz (1991), who studied KIT as a candidate gene in an extended kindred with piebaldism and identiﬁed a missense mutation, Gly664Arg, within the highly conserved intracellular tyrosine kinase domain. This mutation showed complete genetic linkage with the piebald phenotype in this family. Further support came from analyses of two patients with sporadic piebaldism associated with de novo cytogenetic deletions, one of 4q12–q21.1 (Yamamoto et al., 1989) and the other submicroscopic; in both patients, the KIT gene and the adjacent PDGFRA gene were both deleted (Fleischman et al., 1991; Spritz et al., 1992a). Subsequent work has led to the identiﬁcation of at least 37 different point mutations and a number of deletions of the KIT gene in patients with piebaldism and their families (Ezoe et al., 1995; Fleischman, 1992; Fleischman et al., 1996; Giebel and Spritz, 1991; Murakami et al., 2004; Nomura et al., 1998; Richards et al., 2001; Riva et al. 1995; Shears et al., 2004; Spritz et al., 1992b, c, 1993a, b; Syrris et al., 2000, 2002) (Fig. 6.1). About two-thirds of the patients 141

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studied have identiﬁable point mutations, and many of the rest appear to have KIT gene deletions (Ezoe et al., 1995; Fleischman et al., 1991). It is now clear that piebaldism is caused by reduced function of the KIT receptor, resulting in decreased KIT-dependent signaling and consequent abnormal proliferation, survival, or distribution of melanoblasts and melanocytes during embryologic development. Correlation of point mutations of the human KIT gene with the associated clinical phenotypes has proved highly instructive in ascertaining the molecular pathogenic mechanisms underlying piebaldism, as there appears to be a clear and relatively robust genotype–phenotype relationship (Spritz et al. 1992c). As shown in Figure 6.1, KIT point mutations associated with piebaldism can be grouped into four general classes, each associated with phenotypes of differing severity. Thus, a hierarchical paradigm of pathologic human KIT mutations appears to account for a graded series of dominant phenotypes in human piebaldism, correlating reasonably well with predicted effects on KIT-dependent function. The ﬁrst class of KIT gene mutations, all associated with relatively severe piebald phenotypes, consists of missense substitutions located within the highly conserved tyrosine kinase domain. Most of these involve amino acid residues that have been particularly conserved during evolution, and several correspond closely to similar mutations in W mutant mice (reviewed by Morrison-Graham and Takahashi, 1993; Spritz, 1992, 1993; Spritz and Hearing, 1994), underscoring the importance of these sites to the function of the KIT receptor. All these KIT missense substitutions are associated with relatively severe piebald phenotypes. This is a consequence of the fact that the KIT kinase is only activated on dimerization of the receptor, and KIT receptor heterodimers consisting of one normal KIT polypeptide and one abnormal KIT polypeptide are inactive. Thus, patients heterozygous for these “dominantnegative” KIT missense mutant alleles have only one-quarter of the normal amount of KIT receptor dimer and, accordingly, they exhibit relatively severe piebald phenotypes. Even these patients exhibit something of a spectrum of clinical severity, with rare mutations tending to result in extreme phenotypes in which depigmentation may extend to the dorsal aspects of the trunk and head (Fleischman, 1992). The second class of KIT mutations is associated with a relatively wide range of piebald phenotypes, from severe to mild even within an individual family. This group consists of a variety of different nonsense, frameshift, and splice junction mutations, all located distally, within the part of the gene encoding the intracellular portions of the KIT protein that constitutes the tyrosine kinase domain. All these mutations would result in premature termination of translation, truncating the nascent KIT polypeptide distally, within the intracellular tyrosine kinase domain. Clearly, these mutations would abolish the expression of normal KIT polypeptide from this allele. However, such truncated KIT receptors can still bind SLF and even form dimers, dominant negatively inhibiting function of the normal KIT polypeptide (Lev et al., 1992). As both the 142

incompletely translated KIT mRNA and the truncated KIT polypeptide are most likely relatively unstable, even though these truncated KIT polypeptides act dominant negatively, they are present at less than a full dose. Accordingly, this class of mutations presumably reduces KIT function to between onequarter and one-half of normal, accounting for the generally intermediate but highly variable piebald phenotype. The third class of KIT mutations is a group of frameshifts and other “loss of function” mutations that completely eliminate the production of KIT protein by the defective allele. Patients heterozygous for these “loss of function” mutant alleles thus express half the normal amount of KIT receptor, resulting in haploinsufﬁciency for KIT-dependent signal transduction. These KIT mutations are thus associated with relatively mild piebald phenotypes, in which the depigmented patches are usually small, and poliosis is often absent, although affected individuals frequently experience early graying of the hair. In fact, some members of families with KIT mutations of this type have been so mildly affected that the clinical diagnosis was only made subsequent to DNA diagnosis. Interestingly, one of this group of mutations, a four-base deletion (delCAGT) involving codons 250–251, has been observed in at least three unrelated families (Spritz, 1993) and appears to be a recurrent deletion resulting from a direct sequence repetition in this region. The fourth class of KIT mutations constitutes a series of missense substitutions located in the extracellular ligandbinding domain (Fig. 6.1). All these mutations are associated with mild clinical phenotypes (in fact, at least one, Met318Gly, is not pathologic at all). The extracellular domain of the KIT polypeptide consists of ﬁve immunoglobulin-like repeats (Yarden et al., 1987), the ﬁrst three of which bind the SCF ligand (Blechman et al., 1993; Jiang et al., 2000; Lev et al., 1993), and structural redundancy in this region may result in some functional redundancy, thus rendering some amino acid substitutions in this region phenotypically silent. The identiﬁcation of mutations in the KIT gene in human piebaldism opened the door to relatively simple and accurate DNA-based diagnosis and prenatal diagnosis in selected cases. DNA-based molecular diagnosis and even prenatal diagnosis of piebaldism are now routinely accomplished. Sanchez-Martin et al. (2003) have reported heterozygous de novo deletions of the SNAI2 gene (MIM #602150) on chromosome 8q11 in some patients with sporadic piebaldism. SNAI2 encodes Slug, a zinc ﬁnger transcription factor expressed in migratory neural crest cells (Perez-Losada et al., 2002). Although SNAI2 is thus an attractive candidate for a human piebaldism locus, it is puzzling that these same authors have also reported de novo homozygous SNAI2 deletions in patients with sporadic Waardenburg syndrome type 2 (Sanchez-Martin et al., 2002).

Related Disorders KIT was ﬁrst identiﬁed as the active oncogene of the HZ-4 feline sarcoma virus, suggesting that, in contrast to piebaldism, in which KIT expression is reduced, in certain forms of

MELANOBLAST DEVELOPMENT AND ASSOCIATED DISORDERS

human neoplasia or hyperproliferative states, KIT expression might be elevated or dysregulated. Furitsu and coworkers (1993) identiﬁed two KIT gene mutations, Val560Gly and Asp816Val, in the human mast cell leukemia line HMC-1; the Asp816Val substitution constitutively activates the KIT receptor in these cells. Subsequent studies (Buttner et al., 1998; Longley et al., 1996; Nagata et al., 1995) showed that de novo activating mutations of KIT account for most cases of acquired adult mastocytosis and mastocytoma (reviewed by Tsujimura et al., 1997). Activating mutations of KIT have also been found in a small fraction of patients with various types of myeloid leukemias (Beghini et al., 1998; Kimura et al., 1997; Nakata et al., 1995), including about half of adults with core binding factor acute myeloid leukemia (CBFL) (Beghini et al., 2004). Activating mutations of KIT have also been found in a small fraction of gastrointestinal stromal tumors (GIST; MIM #606764) (Hirota et al., 1998; Lasota et al., 1999, 2000; Metaxa-Mariatou et al., 2004; Nakahara et al., 1998; Nishida et al., 1998), but have not been found in most other solid tumors (Chilton-Macneill et al., 2004; Lasota et al., 1999; Lassus et al., 2004; Miettinen et al., 2000; Sihto et al., 2005). In the mouse, a white spotting disorder similar to W is caused by mutations at the Steel (Sl) locus, which encodes SCF, the ligand for the KIT receptor (Lyon and Searle, 1989; Silvers, 1979). The human homolog of the mouse Sl locus gene is called KITLG (for “Kit ligand”). Although aberrant expression of KITLG has been observed in cutaneous mastocytosis (Longley et al., 1993), no abnormalities of the MGF gene have been found in human patients with the piebald phenotype (Ezoe et al., 1995). There may thus be no human homolog to the Steel mutations of mice. Perhaps the function of SCF is redundant in humans, and human KITLG gene mutations thus result in no clinical phenotype, in a clinical phenotype that does not include piebald-like pigmentary anomalies or, alternatively, in humans, KITLG gene mutations might be lethal. Another mouse locus closely related to the Kit/W complex is Patch (Ph), characterized by a dominant white spotting phenotype similar to that of W (Lyon and Searle, 1989; Silvers, 1979). The original Ph mutation was found to comprise a 400kb chromosomal deletion that does not involve the Kit gene at all, but instead deletes the adjacent Pdgfra gene (Nagle et al., 1994; Smith et al., 1991; Stephenson et al., 1991). Although it was therefore initially suggested that white spotting in Ph mice might result from deletion of the Pdgfra gene itself, it was subsequently found that the Ph deletion alters the normal developmental pattern of expression of the cis-linked Kit allele (Duttlinger et al., 1995). Similarly, the mouse rumpwhite (Rw) locus, which is also associated with dominant white spotting (Lyon and Searle, 1989; Silvers, 1979), constitutes a chromosomal inversion that includes Kit (Nagle et al., 1994; Stephenson et al., 1994) and likewise alters the expression of the cis-linked Kit allele during mouse development. Nevertheless, some human GISTs have recently been found to have activating mutations of PDGFRA (Chompret et al.,

2004; Hirota et al., 2003; Lasota et al., 2004; Sakurai et al., 2004; Yamamoto et al., 2004), raising the possibility that the functions of PDGFRA and KIT may overlap to some extent, thus resurrecting the possibility that deletion of Pdgfra might contribute directly to the murine Ph mutant phenotype.

Waardenburg Syndrome Type I (WS1; MIM #193500), Waardenburg Syndrome Type III (WS3; Klein–Waardenburg Syndrome; MIM #148820), and Craniofacial–Deafness–Hand Syndrome (MIM #122880) As discussed in Chapter 29, WS1 is an autosomal dominant disorder characterized clinically by piebald-like hypopigmented patches, heterochromia irides, dystopia canthorum, and sensorineural deafness. WS3 is similar to WS1, but is additionally associated with musculoskeletal abnormalities. Craniofacial–deafness–hand syndrome is similar to WS1, but involves more severe abnormalities of craniofacial development and includes developmental anomalies of the hand. All three of these disorders are now know to be allelic, resulting from mutations of a single gene, PAX3. Elucidation of the molecular bases of WS1 and WS3 was facilitated by studies of a similar disorder of mouse, Splotch (Sp) (reviewed by Pierpont and Erickson, 1993; Spritz, 1993; Spritz and Hearing, 1994). Splotch mice exhibit piebald-like patches of nonpigmented skin and hair (Auerbach, 1954; Epstein et al., 1991, 1993; Goulding et al., 1991, 1993; Moase and Trasler, 1992; Russell, 1947; Vogan et al., 1993), probably due to a reduction in the number of primordial neural crest melanoblasts or to their delayed migration so that melanoblasts fail to reach the affected regions before hair follicles develop (reviewed by Tachibana et al., 2003). The gene for human WS1 was ﬁrst mapped to distal chromosome 2q on the basis of a patient with a chromosomal inversion of 2q35–q37.3 (Ishikiriyama et al., 1989). Subsequent genetic linkage analyses conﬁrmed localization of the WS1 gene to this chromosomal region (Asher et al., 1991; Foy et al., 1990), which corresponds to the location of the murine Splotch locus on mouse chromosome 1, suggesting that human WS1 and mouse Splotch might be homologous. Analysis of the mouse Sp2H mutation showed it to consist of a deletion in the Pax3 gene (Epstein et al., 1991), one of a family of socalled “paired box” (PAX) genes involved in embryologic development (Burri et al., 1989). Two reports reﬁned the localization of the human PAX3 gene to 2q35–q36.2 (Lu-Kuo et al., 1993; Tsukamoto et al., 1992), and Tassabehji et al. (1992) and Baldwin et al. (1992) quickly identiﬁed pathologic PAX3 mutations in patients with WS1. More than 50 different point mutations of the PAX3 gene have since been identiﬁed in patients with Waardenburg syndrome (Baldwin et al., 1994, 1995; Butt et al., 1994; Fortin et al., 1997; Hoth et al., 1993; Morell et al., 1992, 1993; Pardono et al., 2003; Tassabehji et al., 1993, 1994a, 1995; Wildhardt et al., 1996; 143

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Wollnik et al., 2003; Zlotogora et al., 1995), including missense substitutions, nonsense mutations, frameshifts and small in-frame deletions, and splice junction mutations. WS1 is one of the autosomal dominant disorders for which there is evidence of paternal age effect on the occurrence of new mutations (Jones et al., 1975). The PAX3 gene product is a transcription factor that is critical for activating melanoblasts and other cellular elements to proliferate or to begin migration from the neural crest (Stuart and Gruss, 1995). The human PAX3 polypeptide contains four principal structural motifs: a “paired box” domain, a homeobox domain, a conserved octapeptide, and a serinethreonine-proline-rich carboxyl segment. The 128-amino-acid paired box domain, from which the name of the gene is derived, and the 60-amino-acid homeobox domain are both highly conserved amino acid sequence motifs present in a number of mammalian and Drosophila genes involved in controlling segmentation. These motifs form a helix–turn–helix structure and are both thought to mediate DNA binding. Expression of PAX3 is spatially and temporally regulated during development, with localized expression during formation of the embryonic primitive streak (Goulding et al., 1993), during differentiation of the dorsal neuroepithelium (Goulding et al., 1991), during migration of dermomyotomal cells from somites to limb buds (Bober et al., 1994; Franz et al., 1993), and in adult brain (Stoykova and Gruss, 1994). PAX3 acts in concert with SOX10 to trans-activate the promoter of the MITF gene (Bondurand et al., 2000; Watanabe et al., 1998), which encodes one of the key regulators of melanocyte development and melanogenesis (Hornyak et al., 2001; reviewed by Steingrimsson et al., 2004). PAX3 also controls a transcriptional cascade that is involved in regulating skeletal myogenesis (Ridgeway and Skerjanc, 2001). There has been considerable effort to correlate speciﬁc types of mutations of the PAX3 gene with the different clinical subtypes of Waardenburg syndrome, but with only limited success. Clearly, most mutations in the PAX3 gene result in the phenotype of WS1 (Baldwin et al., 1995; Tassabehji et al., 1995). It is apparent that simple loss of function of one allele will result in the WS1 phenotype due to haploinsufﬁciency for the PAX3 gene product. The importance of gene dosage in WS1 is further supported by the observation of several patients with WS1 who have cytogenetic rearrangements that involve PAX3 (Lu-Kuo et al., 1993; Pasteris et al., 1993; Tassabehji et al., 1994a; Tsukamoto et al., 1992; Wu et al., 1993). However, a small fraction of PAX3 mutations do not result in WS1, but instead result in the WS3 (Klein–Waardenburg) phenotype in which features of WS1 are associated with limb anomalies (Hoth et al., 1993; Milunsky et al., 1992; Pasteris et al., 1993; Tassabehji et al., 1995; Tekin et al., 2001). Some patients with WS3 have heterozygous PAX3 mutations, whereas other have homozygous PAX3 mutations (Ayme and Philip, 1995; Wollnik et al., 2003; Zlotogora et al., 1995). Yet other mutations in PAX3 result in “craniofacial–deafness– hand syndrome” (Asher et al., 1996). Thus, WS1, WS3, and craniofacial–deafness–hand syndrome are allelic, and the 144

interplay among functional domains in the PAX3 polypeptide is exceedingly complex (Chalepakis et al., 1994; DeStefano et al., 1998; Fortin et al., 1997), accounting for a remarkable diversity of clinical and developmental phenotypes and patterns of inheritance associated with PAX3 mutations in man. One patient with WS2 was reported to have a mutation in PAX3 (Tassabehji et al., 1993); however, it now seems likely that this patient was misclassiﬁed. No patients with WS2 have mutations in PAX3, and WS2 was found not to be genetically linked to markers from chromosome 2q (Farrer et al., 1992, 1994), indicating that genes responsible for WS2 are not located in this region of the genome. Indeed, it is now clear that at least some patients with WS2 have mutations of the MITF gene, located on chromosome 3p, and others have mutations in the WS2B, WS2C, and WS2D loci, located on chromosomes 1p, 8p, and 8q11 respectively (see below).

Related Disorders Surprisingly, the PAX3 gene also turns out to be rearranged in the t(2;13)(q35;q14) chromosomal translocation frequently observed in cases of alveolar rhabdomyosarcoma (MIM #268220), an aggressive solid tumor of childhood (Barr et al., 1993; Galili et al., 1993). In several such tumors, the chromosome 2 breakpoints were located within the fourth intervening sequence of the PAX3 gene, juxtaposing the upstream PAX3 DNA-binding motifs, including the paired box domain, conserved octapeptide, and homeobox domain, with a speciﬁc “forkhead domain” gene, FOXO1A (FKHR), located on chromosome 13q14.1, which is involved in regulating the growth of a number of different cell types (Medema et al., 2000; Modur et al., 2002; Nakamura et al., 2000). As a result, an abundant novel mRNA is transcribed that encodes a PAX3/FOXO1A fusion protein, which interferes with the normal control of striated muscle cell proliferation (Fredericks et al., 1995; Khan et al., 1999; Lagutine et al., 2002; Relaix et al., 2003; Scheidler et al., 1996; Sublett et al., 1995). Thus, the PAX3 gene appears to lead a double life: inherited loss of function mutations result in Waardenburg syndrome, and acquired gain of function mutations result in alveolar rhabdomyosarcoma, consistent with a role for PAX3 in the developmental regulation of skeletal myogenesis (Ridgeway and Skerjanc, 2001).

Waardenburg Syndrome Type II (WS2; MIM #193500) As discussed in Chapter 36, the phenotype of WS1 is very similar to that of WS1 except that patients with WS2 lack the clinical feature of dystopia canthorum (Arias, 1971; Farrer et al., 1992). Human WS2 can be caused by mutations in at least four different genes, only two of which have thus far been identiﬁed. WS2A (MIM #193510) results from mutations in the MITF gene on chromosome 3p. WS2B (MIM #600193) maps to chromosome 1p and WS2C (MIM #606662) to chro-

MELANOBLAST DEVELOPMENT AND ASSOCIATED DISORDERS

mosome 8p. WS2D (MIM #608890) results from mutations in SNAI2 on chromosome 8q11. Genetic linkage analysis ﬁrst demonstrated that WS2 did not result from mutations in the PAX3 gene (Farrer et al., 1992, 1994), and subsequent studies demonstrated genetic linkage between some (but not all) cases of WS2 (WS2A) and the MITF gene (Hughes et al., 1994), the human homolog of the mouse microphthalmia (mi) locus (Tachibana et al., 1994), located in human chromosome segment 3p12. Pathologic mutations of the human MITF gene have since been identiﬁed in a number of patients with WS2A (Lalwani et al., 1998; Morell et al., 1997; Nobukuni et al., 1996; Smith et al., 2000; Tassabehji et al., 1994b, 1995). Moreover, re-evaluation of a family with “Tietz albinism–deafness syndrome” (Tietz, 1963; MIM #103500) demonstrated that this family segregates a pathologic MITF mutation (Smith et al., 2000). The MITF gene product is a homodimeric transcription factor of the basic helix–loop–helix leucine zipper class (Hemesath et al., 1994; Hodgkinson et al., 1993). In addition to its involvement in regulating melanocyte development (reviewed by Steingrimsson et al., 2004), the MITF protein also appears to play a role in regulating melanocyte function, as it is critical for transcriptional activation of the genes encoding tyrosinase (TYR; Bentley et al., 1994; Tachibana et al., 1996; Yasumoto et al., 1994) and TRP-1 (TYRP1; Yavuzer et al., 1995). Indeed, Morell et al. (1997) showed that the phenotype of WS2A plus ocular albinism (OA) apparently results from “digenic” inheritance, the coincidence of a MITF gene mutation and the R402Q allele of tyrosinase that is associated with reduced enzymatic activity (Tripathi et al., 1991), with resultant downregulation of already reduced tyrosinase catalytic function. The phenotype of the microphthalmia mutant mouse includes hypopigmentation, reduced eye size and cataracts, osteopetrosis, and early onset of deafness, and various different alleles can be either dominant or recessive (Lyon and Searle, 1989; Moore, 1995; Silvers, 1979). Interestingly, although only a modest number of human MITF mutations have been identiﬁed in patients with WS2, one of these is precisely homologous to the mouse mi mutation itself, an inframe deletion of one of a run of four arginines in the basic domain (Tassabehji et al., 1995), and is associated with a somewhat atypical clinical phenotype. Dominant mouse mi alleles operate via dominant negative effects (Hemesath et al., 1994; Steingrimsson et al., 1994), whereas mi mutations that prevent dimerization are recessive. Nevertheless, it is clear that, in humans, WS2A can result from loss-of-function mutations of MITF, which presumably result in haploinsufﬁciency for MITF function. It is intriguing that patients with WS1 resulting from PAX3 mutations and patients with WS2A due to MITF mutations exhibit virtually identical clinical phenotypes except for the absence of dystopia canthorum in WS2. It may be that the embryologic processes responsible for dystopia canthorum are more sensitive to dosage of the PAX3 protein than of the MITF protein. Even within WS1 families with known PAX3

mutations, not all affected individuals exhibit dystopia canthorum, and it has been suggested that variation at some other genetic locus may be responsible for much of this clinical variation in WS1 (Reynolds et al., 1996). Overall, only a relatively small fraction of families with WS2 appears to have mutations in the MITF gene (Hughes et al., 1994; Tassabehji et al., 1995). Lalwani et al. (1994) mapped a second WS2 locus, designated WS2B, to chromosome 1p21–p13.3. Selicorni et al. (2002) assigned WS2C on the basis of a family with a translocation breakpoint involving 8p23. WS2D corresponds to the SNAI2 locus on 8q11 (Sanchez-Martin et al., 2002), with these authors detecting homozygous SNAI2 deletions in two of 38 unrelated WS2 patients who lacked MITF mutations. SNAI2 encodes Slug, a zinc ﬁnger transcription factor expressed in migratory neural crest cells (Perez-Losada et al., 2002). Whereas SNAI2 is thus an attractive Waardenburg syndrome candidate locus, this result is puzzling as these same authors have reported de novo heterozygous SNAI2 deletions in patients with sporadic piebaldism (Sanchez-Martin et al., 2003). Yet another WS locus may reside on chromosome 13q, as van Camp et al. (1995) reported a patient with both WS2 and Hirschsprung disease with a de novo interstitial cytogenetic deletion of chromosome 13q. These same authors also reported a family in which two brothers both exhibited WS2 and Hirschsprung disease, and in which linkage to PAX3 and MITF could be excluded, but linkage to 13q markers could not (see also below).

Waardenburg Syndrome Type IV (WS4; Waardenburg–Shah Syndrome; MIM #277580) As discussed in Chapter 36, WS4 is an autosomal recessive disorder phenotypically similar to WS3, although sometimes with more extreme hypopigmentation (Gross et al., 1995), but without deafness, and involving the additional feature of Hirschsprung disease (HSCR; MIM #142623), the congenital absence of intrinsic ganglion cells of the myenteric and submucosal plexi of the gastrointestinal tract (aganglionic megacolon) (Gross et al., 1995; Shah et al., 1981). Like melanocytes, gut enteric plexus ganglion cells are derived from the neural crest, and thus it is not surprising that there is an association between these phenotypic features. In fact, HSCR has been reported in rare cases of piebaldism, WS1, and WS2; similarly, Hirschsprung disease itself has occasionally been associated with pigmentary anomalies, in both humans and mice. Evidence has begun to emerge for a complex network of interacting genes and proteins that regulate the development of melanocytes and enteric plexus neurons during embryogenesis (Barsh, 1995). Indeed, it is now clear that the WS4 phenotype can result from mutations in genes encoding at least three of these regulatory factors: endothelin-3 (EDN3), the endothelin-B receptor (EDNRB), and the transcription factor SOX10. 145

CHAPTER 6

WS4 Associated with Mutations of the Endothelin-3 Gene (EDN3; MIM #131242) Targeted disruption of the mouse Edn3 gene, encoding endothelin-3, resulted in a recessive phenotype of Hirschsprung disease and spotted coat color (Baynash et al., 1994), similar to the phenotype of human WS4. This laboratory induced mutation is not complemented by the natural mouse mutation, lethal spotting (ls), which was found to result from a point mutation in the Edn3 gene (Baynash et al., 1994). Hood et al. (1989) described a child with sporadic Hirschsprung disease, deafness, and piebald-like white spotting associated with a de novo translocation t(7;20)(q22.1;q13.13); the location of the EDN3 gene at 20q13.2–q13.3 (Arinami et al., 1991; Gopal Rao et al., 1991) suggested that EDN3 may have been disrupted by this translocation. These observations, as well as earlier ﬁndings involving the EDNRB gene, encoding the endothelin-3 receptor (discussed below), made the human EDN3 gene a likely candidate for WS4. Subsequently, two groups identiﬁed homozygous or heterozygous mutations of the EDN3 gene in patients with WS4 (Edery et al., 1996; Hofstra et al., 1996), conﬁrming the homology between human WS4 and murine ls. Mutations of EDN3 may also result in isolated Hirschsprung disease without pigmentary anomalies (Bidaud et al., 1997; Svensson et al., 1999), as well as congenital central hypoventilation syndrome associated with Hirschsprung disease (Bolk et al., 1996).

WS4 Associated with Mutations of the Endothelin Receptor B Gene (EDNRB; MIM #131244); Hirschsprung Disease/Waardenburg Syndrome (HSCR2; MIM #600155); ABCD Syndrome (MIM #600501) Puffenberger and coworkers (1994a) initially mapped a locus associated with recessive “multigenic Hirschsprung’s disease,” frequently associated with white forelock, to chromosome segment 13q22 (Puffenberger et al., 1994a), and van Camp et al. (1995) reported a patient with clinical features of WS2A and a de novo interstitial deletion of chromosome 13q, as well as a family with two siblings with WS2 and Hirschsprung disease. The localization of the EDNRB gene to this same chromosome segment (Puffenberger et al., 1994b), and the fact that both targeted and natural (piebald-lethal; sl) mutations of the mouse Ednrb gene result in a similar recessive phenotype of Hirschsprung disease and spotted coat color (Hosoda et al., 1994) due to a profound deﬁciency of neural crest-derived melanocytes (Pavan and Tilghman, 1994), made EDNRB an obvious candidate for WS4. Two groups (Attie et al., 1995a; Puffenberger et al., 1994b) then identiﬁed mutations of the EDNRB gene, encoding the endothelin B (endothelin-3) receptor, in patients with HSCR2, an autoso146

mal dominant disorder in which Hirschsprung disease is associated with pigmentary anomalies, overlapping the clinical deﬁnition of WS4. Additional dominant EDNRB mutations have since been identiﬁed in other WS4 patients (Syrris et al., 1999). Recessive EDNRB mutations can also result in isolated Hirschsprung disease (Amiel et al., 1996; Kusafuka et al., 1996; Puffenberger et al., 1994a), as well as ABCD syndrome (MIM #600501), characterized by bilateral deafness, albinism, black scalp lock, and retinal depigmentation (Gross et al., 1995; Verheij et al., 2002).

WS4 Associated with Mutations of the SOX10 Transcription Factor Gene (SOX10; MIM #602229); Yemenite Deaf–Blind Hypopigmentation Syndrome (MIM #601706) SOX10 is mutated in the spontaneous dominant megacolon (Dom) mouse (Herbarth et al., 1998; Southard-Smith et al., 1998) and encodes a transcriptional activator related to that encoded by the testis-determining gene SRY. Heterozygous SOX10 mutations have been identiﬁed in a number of patients with WS4 (Kuhlbrodt et al., 1998; Pingault et al., 1998; Southard-Smith et al, 1999), with the so-called WS4 “neurologic variant” (Inoue et al., 1999, 2002; Touraine et al., 2000), and one patient with the “Yemenite deaf–blind hypopigmentation syndrome” (MIM #601706; Bondurand et al., 1999). It is thought that SOX10 mutations that cause pure loss of function may result in WS4, whereas mutations associated with neurological dysfunction may act via dominant-negative mechanisms (Inoue et al., 2002, 2004). SOX10 strongly transactivates the MITF promoter, acting in synergy with PAX3 (Bondurand et al., 2000; Lee et al., 2000; Potterf et al., 2000).

Related Disorders An autosomal dominant form of Hirschsprung disease (MIM #142623) is associated with mutations in the RET protooncogene (Angrist et al., 1995), located at chromosome 10q11.1 (Edery et al., 1994; Romeo et al., 1994), which appears to be a major locus for sporadic Hirschsprung disease (Angrist et al., 1995; Attie et al., 1995b; Bolk Gabriel et al., 2002; Seri et al., 1997). RET encodes a receptor tyrosine kinase related to the torso gene of Drosophila, although its natural ligand is not yet known. RET gene mutations have most frequently been associated with multiple endocrine neoplasia type 2 (MEN2; MIM #171400 and 162300; Eng et al., 1994; Mulligan et al., 1993, 1994), a disorder involving tumors of multiple neural crest-derived endocrinologic tissues and occasional Hirschsprung disease (Eng, 1996; Lyonnet et al., 1993; Mahaffey et al., 1990; Verdy et al., 1982), and with medullary thyroid carcinoma (MTC; MIM #155240; Marsh et al., 1996). However, RET mutations have not yet been associated with developmental pigmentary anomalies in humans.

MELANOBLAST DEVELOPMENT AND ASSOCIATED DISORDERS

Aganglionic megacolon is a relatively frequent occurrence in patients with Down syndrome (Garver et al., 1985; Passarge, 1967), and evidence has been presented for a “modiﬁer locus” on human chromosome 21q22 that affects expression of Hirschsprung disease in patients with EDNRB gene mutations (Puffenberger et al., 1994a). Up to six genetic modiﬁer loci may modulate the severity of white spotting associated with homozygosity for Ednrb mutations in piebald (s) mice (Pavan et al., 1995). Whereas some of these modiﬁers may be known genes, such as W and Sl, others may well be novel genes that may themselves be associated with developmental mutations that affect pigmentation. It is of some interest, for example, that the human homolog of the Drosophila white gene is located at 21q22.3 (Chen et al., 1996), although this gene has not thus far been associated with pigmentary abnormalities in humans.

X-linked Albinism–Deafness Syndrome (ADFN; MIM #300700) ADFN is an X-linked recessive disorder characterized by extreme piebald-like hypopigmentation and deafness affecting only males in a large Israeli kindred (Margolis, 1962; Ziprkowski et al., 1962). Genetic linkage and somatic cell hybridization studies have localized the gene to Xq24–q26 (Shiloh et al., 1990); however, as of this writing, the gene for this disorder had not yet been identiﬁed.

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The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

7

Biogenesis of Melanosomes Raymond E. Boissy, Marjan Huizing, and William A. Gahl

Introduction The major product of the melanocyte is the specialized organelle called a pigment granule, or melanosome. As discussed in the preceding chapters, the cutaneous melanocyte synthesizes this organelle for eventual transfer to neighboring keratinocytes. In contrast, noncutaneous melanocytes retain their quota of pigmented melanosomes. This chapter addresses the origin of this specialized organelle, the processing and transport of gene products destined to occupy this organelle, and the melanosome’s maturation to a specialized microenvironment.

Origin of the Stage I Premelanosome During the early part of the twentieth century, pigmented tissue such as skin and eyes was shown to have a dark granular morphology when examined by light microscopy. This prompted biologists to speculate that pigment granules could originate from: (1) nuclei via extrusion; (2) mitochondria via conversion; or (3) the Golgi apparatus via production (Barnicot and Birbeck, 1958; Guttes, 1953). When investigators began exploring pigmented tissue using electron microscopy in the latter half of the twentieth century, it was initially concluded that pigmented organelles formed from colorless, thick-walled, hollow ellipsoids in which a ﬁbrous matrix was laid down and on to which melanin was subsequently deposited (Barnicot et al., 1955; Birbeck et al., 1956). These observations resulted in discarding the hypothesis that pigment granules originate from a mitochondrial or nuclear origin. At this time, the various ultrastructural proﬁles of pigment granules were collaterally termed “melanosomes” and staged into morphological categories suggesting a developmental process (Birbeck, 1963; Moyer, 1961; Seiji et al., 1961, 1963). To this day, these stages are routinely used. They consist of: Stage I (also termed “premelanosome”) — a relatively spherical organelle with matrix material (i.e. ﬁlaments) beginning to assemble; Stage II — an oval-shaped organelle containing an organized internal matrix without the appearance of melanin; Stage III — an organelle exhibiting deposition of melanin on the matrix; and Stage IV — an organelle completely ﬁlled with melanin (Fig. 7.1). During the initial ultrastructural analyses of melanosomes, Moyer (1961) identiﬁed Stage I melanosomes associated with membranes of the endoplasmic reticulum (ER). Speciﬁcally, Moyer stated that, in

embryonic retinal pigmented epithelium, “one often ﬁnds vesicular dilations in the ER that are approximately the same size as stage I granules and they usually appear empty but occasionally show traces of very thin ﬁbers and small vesicles in the lumen” (Moyer, 1961). This suggested that the premelanosome originated from the ER. Many subsequent authors agreed with this idea (Eppig and Dumont, 1972; Ide, 1972; Maul, 1969; Maul and Brumbaugh, 1971; Stanka, 1971; Stanka et al., 1981). With the advent of DOPA histochemistry, evidence for a non-Golgi origin of melanosomes was strengthened. DOPA incubation is a technique in which this exogenously supplied tyrosinase substrate is converted into melanin at cellular sites where functional tyrosinase resides. As a consequence of this experimental procedure, many trans-Golgi elements and perinuclear melanosomes increased in reaction product (Fig. 7.2). However, many premelanosome organelles in the vicinity of the Golgi remained negative for the reaction product, suggesting that melanosomes did not originate directly from the Golgi (Maul, 1969; Maul and Brumbaugh, 1971). In addition, many 50-nm coated vesicles appeared to fuse with the DOPA-negative premelanosomes, suggesting that the tyrosinase enzyme was trafﬁcked to the preformed, recipient premelanosome. This “bipartite” hypothesis of melanosome biogenesis was substantiated by Turner et al. (1975), who demonstrated that premelanosome structures were derived from the ER in the goldﬁsh. In addition, Turner et al. (1975) demonstrated that DOPA-positive vesicles fused with the premelanosome, inverted, and reformed within the lumen. Vesicles within premelanosomes were originally described by Tousimis (1963), and these vesicles sometimes “opened directly into the cytoplasm.” This was subsequently reported by Durrer and Villiger (1967). These vesicles were further explored by Jimbow et al. (Jimbow and Fitzpatrick, 1974; Jimbow and Kukita, 1970; Jimbow et al., 1979) and proposed to be inverted vesicles trafﬁcking tyrosinase. By the end of the 1990s, conventional dogma held that premelanosomes formed from the ER and that enzymes and regulatory proteins were subsequently trafﬁcked to them from the trans-Golgi in 50-nm clathrin-coated vesicles. However, this concept is currently being challenged in light of recent research focusing on a molecule called Silver/Pmel17. Silver/Pmel17 (also known as Pmel17, gp100, ME20, RPE1, and the silver locus protein) is a type 1 transmembrane glycoprotein that localizes to the matrix of Stage I and II melanosomes (Kobayashi et al., 1994; Lee et al., 1996; Raposo 155

CHAPTER 7

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II G CE III

IV

CE

I

M

IV G

I

III

II G

M

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Fig. 7.1. Perinuclear area of normal human melanocytes from a lightly pigmented Caucasian skin demonstrating melanosomes in various stages of maturation. Melanosomes in the vicinity of the Golgi apparatus (G) exist as Stage I premelanosomes (I) that are more oval in shape containing ﬂoccular material and short irregular melanoﬁlaments, Stage II melanosomes (II) that are more elongated containing an organized, scaffold-like matrix of melanoﬁlaments, Stage III melanosomes (III) with melanin being sequentially deposited on the melanoﬁlament matrix, and Stage IV melanosomes (IV) with melanin completely ﬁlling the organelle and obscuring the matrix. This melanocyte is from a lightly pigmented Caucasian individual and consists of Stage II and III melanosomes predominantly with relatively few Stage IV melanosomes. CE, coated endosome; N, nucleus; M, mitochondria. Bar = 1.0 mm.

et al., 2001). Silver/Pmel17 appears to be the initial melanocyte-speciﬁc molecular marker for melanosomes. Consistent with the hypothesis that the pre-/stage I melanosome originates from the ER, it was demonstrated in mouse melanocytes that Silver/Pmel17 lacks Golgi-modiﬁed N-linked glycans (Kobayashi et al., 1994) and coexists with ER-resident proteins (e.g. calreticulin, calnexin, ribophorin) in puriﬁed premelanosomes (Basrur et al., 2003; Kobayashi et al., 1994; Kushimoto et al., 2001). However, other data suggest that premelanosomes arise from compartments of the late secretory (post-Golgi) and/or endocytic pathways. Golgi-processed, glycosylated forms of Silver/Pmel17 have been identiﬁed in human premelanosomes (Berson et al., 2001; Raposo et al., 2001), suggesting that the delivery of this molecule to a structure designating it as a melanosome occurs post Golgi. Detailed ultrastructural studies (Raposo et al., 2001) showed that Silver/Pmel17 was enriched in coated, multivesicular, endosomal compartments containing protruding

156

I

Fig. 7.2. Perinuclear area of normal human melanocytes from a lightly pigmented Caucasian skin demonstrating localization of tyrosinase by l-DOPA histochemistry. DOPA reaction product indicating cellular sites of catalytically functional tyrosinase appears in the trans-Golgi network (G) including the trans-most cisternae (arrows), coated vesicles budding from these cisternae (arrowheads), and vesicles migrating from the Golgi apparatus (arrowheads with asterisk). Minimal reaction product exists in some Stage I premelanosomes (I), indicating incorporation of tyrosinase into these organelles. After DOPA histochemistry, this lightly pigmented, Caucasian melanocyte consists of Stage IV melanosomes predominantly with relatively few Stage II and III melanosomes. CE, coated endosome; N, nucleus; M, mitochondria. Bar = 1.0 mm.

smooth tubules. This compartment was shown to evolve into a multivesicular endosomal intermediate, the coated endosome, proposed to represent the premelanosome (Fig. 7.3) (Raposo et al., 2001; Raposo and Marks, 2002). The coated endosome contains large clathrin lattices on the cytoplasmic side (Fig. 7.3), which are thought to function in sorting premelanosome-bound cargo, including Silver/Pmel17, away from other endocytic cargo bound for lysosomes. Currently, unidentiﬁed chaperones likely facilitate this sorting, perhaps by recognizing speciﬁc domains of Silver/Pmel17 and other premelanosomal components. One candidate chaperone in this sorting process is adaptor protein-3 (AP3). Coated endosomes are also found in nonpigmented cells and are endosomal precursors of multivesicular bodies (Sachse et al., 2002). Proteomic analysis revealed the presence of Silver/Pmel17 in the premelanosome, but tyrosinase and other melanosomal proteins were also detectable (Kushimoto et al., 2001). However, the highly proteolytic environment in these vesicles appears to degrade or partially cleave these proteins and impair their enzymatic activities. Silver/Pmel17 also undergoes

BIOGENESIS OF MELANOSOMES

CE I

CE

? A

CE CE I I

B

C

Fig. 7.3. Proﬁles of coated endosomes/premelanosomes within the perinuclear area of normal human melanocytes. Perinuclear areas of normal human melanocytes without (A and B) or with (C) DOPA histochemistry demonstrate coated, multivesicular endosomal compartments (CE) proposed by Marks and Raposo (Raposo et al., 2001; Raposo and Marks, 2002) as precursor organelles developing into premelanosomes (I). These structures are characterized by focal, clathrin lattices (arrowheads) present on the cytoplasmic face. Proﬁles ultrastructurally resembling intermediates between the coated, multivesicular endosome and the premelanosome with melanoﬁlaments undergoing assembly can be observed occasionally (?). Bar = 1.0 mm.

cleavage in this environment by a furin-like proprotein convertase (Berson et al., 2003). Properly cleaved Silver/Pmel17 has a key function in restructuring the premelanosomal matrix into ﬁbrous striations, leading to a Stage II melanosome (Berson et al., 2003). In conclusion, it remains unsubstantiated whether the organelle that becomes a premelanosome originates from the ER and/or cis-Golgi or as a specialization of the endosomal compartment. In addition, the discrepancy regarding the presence of tyrosinase and Tyrp1 in early Stage I melanosomes is not completely resolved.

Development of the Melanosome Regardless of the actual origin of the premelanosome, it eventually becomes a distinct organelle with a specialized microenvironment to which a battery of enzymes and regulatory proteins are trafﬁcked to facilitate the conversion of tyrosine to melanins. The major group of enzymes and regulatory proteins that eventually reach the melanosome are members of the tyrosinase gene family [tyrosinase-related proteins

(TRPs)], i.e. tyrosinase, tyrosinase-related protein-1 (Tyrp1), and dopachrome tautomerase/tyrosinase-related protein-2 (Dct/Tyrp2). These Type 1 membrane glycoproteins are transcribed at the rough endoplasmic reticulum (RER), shuttled through the Golgi apparatus for carbohydrate maturation, and ultimately transported to premelanosomes in the vicinity of the trans-Golgi network (TGN) via 50-nm coated vesicles. The initial processing of TRPs has been thoroughly explored in recent years. All three TRPs exhibit similar initial processing events, but distinct differences exist. Tyrosinase is initially translated as a 60-kDa polypeptide that is heavily glycosylated in the RER to a 70- to 74-kDa molecular weight form (Muller et al., 1988; Ujvari et al., 2001). Tyrosinase contains up to seven Asn-X-Ser/Thr sequons, and Glc3Man9GlcNAc2 glycans are added to six or seven of them (Branza-Nichita et al., 2000a; Ujvari et al., 2001). The proper amount of glycosylation, the appropriate folding into a tertiary structure, and the acquisition of copper are regulated by glycosylation enzymes and RER chaperone molecules such as calnexin (Branza-Nichita et al., 1999; Toyofuku et al., 1999; Wang and Androlewicz, 2000). Calnexin is a lectin involved in ER quality control that acts by driving the incompletely folded nascent chains into glucosylation/deglycosylation cycles (Ellgaard and Helenius, 2003; Kowarik et al., 2002; Zapun et al., 1997). Mutations in tyrosinase, either experimentally induced (Branza-Nichita et al., 2000b) or occurring naturally in patients with oculocutaneous albinism type 1 (Halaban et al., 2000; Park et al., 1993; Spritz et al., 1990), indicate that aberrations in N-glycosylation or conformational structure result in the degradation of the abnormal molecule via an ER-associated and/or proteasome mechanism (Berson et al., 2000; Halaban et al., 1997; Mosse et al., 1998). The mechanism of ER-associated protein degradation has been reviewed recently (Jarosch et al., 2002). Copper binding, speciﬁcally at the CuB site (Olivares et al., 2003), also appears to facilitate the initial processing of tyrosinase into a form capable of being transported out of the RER. In addition, DOPA, tyrosine (Halaban et al., 2001), and Tyrp1 (Kobayashi et al., 1998) apparently facilitate the maturation of tyrosinase in the RER. The initial synthesis of Tyrp1 and Dct (Tyrp2) in the RER is similar to that of tyrosinase. Tyrp1 is also a glycoprotein containing six sites with glycosylation sequons (BranzaNichita et al., 2000a; Xu et al., 2001). It is initially translated as a 55-kDa polypeptide, ultimately matures to a 75-kDa glycoprotein (Vijayasaradhi et al., 1991), and constitutes the predominant tyrosinase-related protein synthesized by the melanocyte (Vijayasaradhi et al., 1990). Tyrp1 undergoes a more rapid carbohydrate processing than tyrosinase (Negroiu et al., 1999a; Vijayasaradhi et al., 1991), i.e. ranging in duration from 2–3 hours vs. 3–6 hours respectively (Halaban et al., 1997; Negroiu et al., 1999a; Vijayasaradhi et al., 1995). This range varies with cell line but is primarily related to ER residency time. Treatment of melanoma cells with glycosylation inhibitors does result in a modiﬁed conformation

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(Negroiu et al., 2000). However, Tyrp1 is less sensitive to inhibitors of N-glycan processing than is tyrosinase (Negroiu et al., 1999b). Structural analyses of N-linked glycans for tyrosinase and Tyrp1 demonstrate signiﬁcant differences in the extent and complexity of carbohydrate processing, indicating that the two molecules have different accessibility to the same processing milieu, probably because of differences in their three-dimensional structures (Negroiu et al., 1999b). Alterations in Tyrp1 that affect either glycosylation (Toyofuku et al., 2001) or amino acid sequence at the cytoplasmic tail (Xu et al., 1998) result in retention/degradation in the ER. Dct (Tyrp2) is also glycosylated in the ER and shares conserved potential N-glycosylation sites with tyrosinase and Tyrp1 (Xu et al., 2001). Processing of N-glycan in the ER is mediated by calnexin, which also facilitates the formation of stabilizing disulﬁde bridges (Negroiu et al., 2003). Upon proper initial synthesis, the precursor TRPs are trafﬁcked out of the ER to the Golgi, primarily via vesicles directed by COPII coat complexes (Antonny and Schekman, 2001; Duden, 2003; Murshid and Presley, 2004). TRPs are ultimately shuttled through the Golgi apparatus where sequential modiﬁcations of the sugar residues occur. This shuttling, at least for tyrosinase, appears to be regulated in part by glycosphingolipids. A mouse melanoma cell line deﬁcient in the synthesis of glycosphingolipid demonstrates retention of tyrosinase, but not Tyrp1, in the cisternae of the Golgi apparatus (Sprong et al., 2001). Upon reaching the trans-most cisternae of the Golgi, the mature tyrosinase is a catalytically functional enzyme, as demonstrated by DOPA histochemistry (Fig. 7.2) (Maul, 1969; Maul and Brumbaugh, 1971). Interestingly, this catalytically active enzyme is sequestered from its substrate until deposited into the premelanosome. Targeting sequences on the carboxyl ends of the TRP molecules are essential for accurate targeting to the melanosome (Table 7.1). Tyrosinase contains a dileucine-based motif with ﬂanking regulatory amino acids (two leucine or isoleucine residues with an acidic residue four or ﬁve positions upstream) that interacts with adaptin-3 (AP3)

to facilitate its shipment to the melanosome (Calvo et al., 1999; Honing et al., 1998; Simmen et al., 1999). In the absence of this interaction, as in AP3 mutants (human HPS2, pearl, and mocha mice), tyrosinase does not reach the premelanosome efﬁciently (Huizing et al., 2001a). Tyrp1 also contains carboxy-terminal dileucine sorting signals that mediate its trafﬁcking. Truncated Tyrp1 molecules are erroneously diverted to the plasma membrane (Vijayasaradhi et al., 1995). There is a requisite dileucine motif that exists in the carboxy region, but it is not utilized by AP3 for trafﬁcking (Huizing et al., 2001a) because it lacks the correct adjoining/regulatory amino acid sequence. Instead, Tyrp1 may exit the trans-Golgi network (TGN) via an adaptor complex-1 (AP1)-mediated mechanism (Raposo et al., 2001). Recently, Vijayasaradhi (Liu et al., 2001a) identiﬁed by yeast twohybrid screening of a melanocyte cDNA library the interaction of Tyrp1 with GIPC, a member of a large family of PDZ domain proteins that recognize the sequence motif Ser/ThrXaa-Val on putative interactive/cargo proteins (Songyang et al., 1997). Tyrp1 expresses carboxyl-terminal Ser-Val-Val residues that interact transiently with GIPC after synthesis (Liu et al., 2001a). Interestingly, GIPC cofractionates with adaptin 1 (De Vries et al., 1998), suggesting that Tyrp1 utilizes AP1 as opposed to AP3 in its trafﬁcking. While the speciﬁc adaptor complexes utilized for tyrosinase and Tryp1 trafﬁcking differ, a dileucine motif functions in melanosomal sorting of both these melanogenic proteins. Tyrosine-based sorting signals (YXX F, Y = tyrosine, X = any amino acid, F = bulky hydrophobic amino acid), present in tyrosinase, Tyrp1 and Dct/Tyrp2, may also play a role in the sorting process (Simmen et al., 1999). This raises the possibility that melanosomal proteins follow alternative paths to the melanosome by utilizing tyrosine-based sorting signals and AP1 recognition for sorting at the TGN, and AP2-mediated sorting for internalization from the plasma membrane. Another possibility is that tyrosine-based signals are utilized to retrieve mistargeted melanosomal proteins. Melanosomal proteins may also utilize signals different from tyrosine and

Table 7.1. Putative sorting signals in the cytoplasmic domains of proteins targeted to the melanosome. Protein

Human (murine) disorder

Targeting motif

Tyrosinase Tyrp1 DCT/Tyrp2 Pmel17 p MART1/MLANA OA1

OCA1 (albino, platinum) OCA3 (brown) Unknown (slaty) Unknown (silver) OCA2 (pink-eye dilute) Unknown (unknown) OA1 (Moa1)

TMD-X9-EEKQPLLMEKEDYHSLYQSHL-cooh TMD-X8-DEANQPLLT-X8-EYEKLQNPNQSVV-cooh TMD-RRLRKGYTPLMETHLSSKRYTEEA-cooh TMD-X25-PRIFCSCPIGENSPLLSGQQV-cooh nh2-X90-KEDTPLLW-X77-TMD7-VV-cooh TMD-X6-GYRALMD-X43-AYEKLSAEQSPPPYSP-cooh TMD-X114-cooh

Bold highlighted gray, dileucine-based sorting signal (two leucine or isoleucine residues with an acidic residue 4 or 5 positions upstream). Highlighted black, tyrosine-based sorting signals (YXXF, Y = tyrosine, X = any amino acid, F, bulky amino acid) (Calvo et al., 1999; Honing et al., 1998; Simmen et al., 1999). Not all the signals shown in this table have been experimentally demonstrated to be important for sorting; -cooh, carboxyl-terminus; nh2-, aminoterminus; TMD = transmembrane domain, OCA = oculocutaneous albinism, OA = ocular albinism. See text for details.

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BIOGENESIS OF MELANOSOMES

dileucine signals, as indicated by the absence of dileucine motifs from Dct/Tyrp2 and the OA1 protein. Dct/Tyrp2 processing exhibits post-translational modiﬁcations that suggest transcription in the RER followed by maturation in the Golgi (Tsukamoto et al., 1992). Some data indicate that Dct/Tyrp2 is predominantly maintained in the TGN (Negroiu et al., 2003) and is less prevalent in late melanosomes. Unlike tyrosinase and Tyrp1, Dct/Tyrp2 does not have a dileucine motif, but it does have a tyrosine-based signal (Table 7.1) (Jackson et al., 1992), resembling LAMP-1 and MART-1 (De Maziere et al., 2002; Dell’Angelica et al., 2000a). This suggests that Dct’s trafﬁcking route/mechanism is distinct from that of tyrosinase and Tyrp1. It is uncertain whether TRPs trafﬁc directly from the Golgi to the melanosome. Tyrosinase and Tyrp1, for example, may transit through a late endosome/lysosome compartment en route to the melanosome (Huizing et al., 2001a; Jimbow et al., 1997; Orlow et al., 1993). Another melanocyte-speciﬁc protein related to melanosome biogenesis is the product of the ocular albinism type 1 (OA1) gene. OA1 is an X-linked recessive disorder in which giant macromelanosomes exist predominantly within ocular but also within cutaneous melanocytes (Garner and Jay, 1980; O’Donnell et al., 1976). The OA1 gene encodes a novel integral membrane protein of seven transmembrane domains with homologies to G protein-coupled receptors (Bassi et al., 1995; Schiafﬁno et al., 1999). Initial localization studies using immunoelectron microscopy suggested that the OA1 protein localizes in part to melanosomes (Schiafﬁno et al., 1999). However, recent studies utilizing immunocytochemistry, cellular fractionation, and transfection of GFP-OA1 analysis has demonstrated that the OA1 protein is predominately localized to a late endosome/lysosome compartment and is largely absent from mature melanosomes (Samaraweer et al., 2001; Shen et al., 2001). This suggests that the OA1 protein is positioned to facilitate in part the trafﬁcking of melanosomedistinct proteins through the intermediate compartment en route to the melanosome or to direct the endolysomal compartment into a melanosome. Loss of function of the OA1 protein may result in accumulation of melanogenic enzymes in the late endosome/lysosome and divert this organelle into the unique macromelanosome, as proposed by Shen et al. (2001). Chemically, melanins have been divided into the black/ brown eumelanins and the red/yellow, sulfur-containing pheomelanins. Correlative morphologic features distinguish melanosomes that exist in tissues predominantly eumelanotic vs. pheomelanotic (Hearing et al., 1973; Jimbow et al., 1979). Eumelanosomes are relatively oval organelles, in which the melanoﬁlaments organize into a regularly registered matrix. In contrast, pheomelanosomes are relatively spherical organelles, in which the melanoﬁlaments exist in short, irregularly arranged proﬁles. As melanin synthesis progresses, eumelanosomes acquire a uniform pigmentary pattern whereas pheomelanosomes exhibit an uneven deposition of pigment. It is currently unknown what regulates the unique features of these two types of melanosomes. Interestingly,

expression of Tyrp1, Dct/Tyrp2, and the silver protein (Pmel17) is not present in tissue undergoing pheomelanogenesis (Kobayashi et al., 1995), suggesting that the presence of these melanosome-residing, transmembrane proteins may impart some structural integrity to the eumelanosome.

Chaperoning of Tyrosinase Family Members to the Premelanosome Numerous chaperone proteins, responsible for the accurate trafﬁcking and sorting of the tyrosinase family member proteins from the Golgi apparatus to the premelanosomes, are being identiﬁed. Many of these chaperones are encoded by genes that result in human and murine pigmentation disorders, such as Hermansky–Pudlak syndrome (HPS), Chediak–Higashi syndrome (CHS), and Griscelli syndrome (GS) (Huizing et al., 2000). These disorders of lysosomerelated organelle biogenesis typically affect melanosomes, platelet dense granules, and lysosomes; patients consequently show symptoms of hypopigmentation, prolonged bleeding, and other abnormalities of the lung, gut, and immune system (Clark and Grifﬁths, 2003; Dell’Angelica et al., 2000b; Huizing et al., 2000; Shiﬂett et al., 2002). HPS patients have oculocutaneous albinism, absent platelet dense bodies, and, sometimes, pulmonary ﬁbrosis and granulomatous colitis (Huizing et al., 2000, 2002a; Huizing and Gahl, 2002). CHS patients manifest giant intracellular inclusions in leukocytes and platelets; without a bone marrow transplant, they die of infections in the ﬁrst decade of life (Barak and Nir, 1987; Huizing et al., 2001b; Introne et al., 1999; Shiﬂett et al., 2002; Ward et al., 2002). GS patients also have immunological problems and may manifest neurological disorders as well (Anikster et al., 2002; Griscelli et al., 1978; Sanal et al., 2002). Most CHS and GS patients who do not undergo a bone marrow transplant eventually experience a fatal accelerated phase of lymphocytic proliferation. Identiﬁcation of the chaperone-encoding genes involved in the above disorders can help us to unravel the intertwined biochemical machinery active in melanosome biogenesis. The best understood chaperone in this set is adaptor protein-3 (AP3). This ubiquitously expressed heterotetrameric complex binds clathrin to provide structure to newly forming vesicles and bind dileucine sorting motifs in the cytoplasmic tails of cargo proteins destined to be part of the nascent organelles (Dell’Angelica et al., 1997a, b; Simpson et al., 1997). Defects in a speciﬁc AP3 subunit, b3A (AP3B1), have been identiﬁed in human HP syndrome type 2 or HPS-2 (Clark et al., 2003; Dell’Angelica et al., 1999; Huizing et al., 2002b; Shotelersuk et al., 2000). In addition, AP3 subunit defects are responsible for the pearl (Ap3b1) (Feng et al., 1999) and mocha (Ap3d) (Kantheti et al., 1998) mouse coat color mutants and for pigmentary eye defects in the Drosophila melanogaster mutants ruby (Kretzschmar et al., 2000) and garnet (Ooi et al., 1997). Current dogma suggests that AP3 mediates melanosomal cargo sorting and vesicle budding at the TGN, but AP3 may 159

CHAPTER 7

also operate at other intracellular locations. AP3 could function in the retrieval of tyrosinase, and possibly other melanosomal proteins, from endosomal membranes, with subsequent placement in vesicles destined to become mature melanosomes. AP3 could also retrieve tyrosinase from melanosomal membranes in order to maintain homeostatic levels of the enzyme (Peden et al., 2004; Raposo and Marks, 2002). In AP3-deﬁcient melanocytes, tyrosinase accumulates in multivesicular structures similar in appearance to the coated endosome (Huizing et al., 2001a). This could be due to missorting at the TGN or to failure to retrieve tyrosinase from endosomes. In contrast to tyrosinase, Tyrp1 localization does not appear to be affected by AP3 deﬁciency (Huizing et al., 2001a). In fact, colocalization of Tyrp1 with AP1 in melanocytes suggests a role for AP1 in Tyrp1 transport to melanosomes (Raposo et al., 2001). A novel PDZ domain protein, GIPC, might also have a role in Tyrp1 sorting to melanosomes (Liu et al., 2001b). HP syndrome exhibits locus heterogeneity, and genes associated with human HPS subtypes 1, 3, 4, 5, 6, and 7 (and their respective mouse orthologs pale ear, cocoa, light ear, ruby-eye 2, ruby eye, and sandy), all code for proteins that contain no recognizable motifs or homologies that could predict their biochemical function (Huizing et al., 2002a; Huizing and Gahl, 2002). However, some of these proteins can associate with each other into distinct multisubunit complexes, namely BLOC-1, BLOC-2, and BLOC-3 (biogenesis of lysosomerelated organelles complexes), which function to form and trafﬁc intracellular vesicles (Di Pietro et al., 2004; FalconPerez et al., 2002; Gautam et al., 2004; Li et al., 2003; Martina et al., 2003; Nazarian et al., 2003; Starcevic and Dell’Angelica, 2004) (Fig. 7.4).

Snapin

indin Dysb

AP3

BLOC-2

Table 7.2. Human and mouse genes causing HPS and related disorders. Complex

Human gene

Human disorder

Mouse mutant

AP3

AP3B1 (b3A) AP3D1 (d) AP3S1 (s3A) AP3M1 (m3A)

HPS-2 – – –

pearl mocha – –

BLOC-1

HPS7/Dysbindin Pallidin Muted Cappuccino BLOS1 BLOS2 BLOS3 Snapin

HPS-7 – – – – – – –

sandy pallid muted cappuccino

BLOC-2

HPS3 HPS5 HPS6

HPS-3 HPS-5 HPS-6

cocoa ruby-eye 2 ruby eye

BLOC-3

HPS1 HPS4

HPS-1 HPS-4

pale ear light ear

Others

RGGTA Rab38 VPS33A MYO5A RAB27A Melanophilin

– – – GS1 GS2 GS3

gunmetal chocolate buff dilute ashen leaden

LYST

CHS

beige

Pallidin

Muted

BLOC-1

BLOC-3

Fig. 7.4. Assembly of tyrosinase-related protein chaperone complexes. Proposed assembly structures of adaptor complex 3 (AP3) and biogenesis of lysosome-related organelles complexes (BLOCs). ?, undeﬁned subunit(s).

160

BLOC-1 consists of the palladin, muted, cappuccino, HPS7/dysbindin proteins, and the recently identiﬁed novel subunits BLOS1, BLOS2, BLOS3, and Snapin (Falcon-Perez et al., 2002; Li et al., 2003; Martina et al., 2003; Starcevic and Dell’Angelica, 2004). The genes encoding pallidin, muted, cappuccino, dysbindin, and BLOS3 are mutated in the HPS mouse strains pallid, muted, cappuccino, sandy, and reduced pigmentation respectively (Table 7.2) (Ciciotte et al., 2003; Huang et al., 1999; Li et al., 2003; Starcevic and Dell’Angelica, 2004; Zhang et al., 2002). These mouse strains exhibit the most severe coat color dilution phenotypes among all mouse HPS models (Li et al., 2004). Except for one patient with a dysbindin/HPS7 defect (Li et al., 2003), no mutations in any of the other BLOC-1 subunits have been identiﬁed in human HPS patients. The molecular function of BLOC-1 remains unknown, but indications from its potential binding partners suggest that BLOC-1 operates in the regulation of SNARE-mediated membrane fusion. SNAREs are soluble N-ethylmaleimide-sensitive factor attachment protein receptors involved in vesicle membrane fusion. Palladin is able to interact with syntaxin 13, a SNARE family member localized to early endosomes and involved in membrane fusion at early stages of the endocytic pathway (Huang et al., 1999; Prekeris et al., 1998), and

reduced pigment

BIOGENESIS OF MELANOSOMES

Snapin interacts with SNAP-25/23, two closely related SNARE proteins implicated in membrane fusion events (Buxton et al., 2003; Ilardi et al., 1999; Starcevic and Dell’Angelica, 2004). Dysbindin is a binding partner of alpha and beta dystrobrevins, i.e. dystrophin-related proteins that are part of large dystrophin-associated, membrane-spanning complexes functioning in membrane stability and signaling events (Benson et al., 2001). BLOC-1 associates with F-actin (FalconPerez et al., 2002), supporting a role for the complex in promoting vesicle movement in the periphery of cells. BLOC-2, a stable complex containing the HPS3, HPS5, and HPS6 proteins, exists in both human and mouse (Table 7.2) (Di Pietro et al., 2004; Gautam et al., 2004). Clinically, HPS3, HPS-5, and HPS-6 are relatively mild forms of the disease (Anikster et al., 2001; Huizing et al., 2001c, 2004; Tsilou et al., 2004), with no reported involvement of pulmonary ﬁbrosis, suggesting a common biological basis underlying the pathogenesis of these three HPS subtypes. A regulatory role for BLOC-2 in the secretion of lysosomes and related organelles has been suggested (Di Pietro et al., 2004), based on reduced secretion of lysosomal enzymes by kidney and platelets from both ruby eye (HPS-6) and ruby-eye 2 (HPS-5) mice (Swank et al., 1998). In addition, an increase in the frequency and duration of transient fusion events in ruby-eye mast cells during stimulation implicates a function for the ruby-eye gene product in regulating closure of the cell fusion pores (Oberhauser and Fernandez, 1996). These pores connect the lumen of a secretory vesicle with the extracellular environment during exocytosis. However, ﬁbroblasts from cocoa (HPS-3) and ruby-eye (HPS-6) mice showed normal secretory levels of the enzyme b-hexosaminidase, indicating that BLOC2 is not critical for secretion of this lysosomal enzyme (Di Pietro et al., 2004). Recent studies in ﬁbroblasts have shown that HPS3 localizes to clathrin-coated vesicles in the perinuclear area and that HPS3 contains a clathrin-binding domain at residues 172–176 (LLDFE), which is essential for the accurate localization of LAMP1 (Helip-Wooley et al., 2004). Studies on human HPS-3 melanocytes indicate that melanosome-targeted cargos (tyrosinase, Tyrp1, and Tyrp2) do not become incorporated efﬁciently into melanosomes in the perinuclear region but are retained in 50-nm vesicles that extend into the periphery of the HPS-3 melanocyte. Therefore, BLOC-2 and its association with clathrin may be necessary for the efﬁcient incorporation of melanogenic cargo targeted to the melanosome (Boissy et al., 2004). BLOC-3 contains HPS1 and HPS4 (Table 7.2) (Chiang et al., 2003; Martina et al., 2003; Nazarian et al., 2003), which explains the phenotypic similarity of HPS-1 and HPS4 patients. In addition to oculocutaneous albinism and prolonged bleeding, present in all HPS subtypes, HPS-1 and HPS-4 patients are at high risk of developing progressive pulmonary ﬁbrosis and granulomatous colitis (Anderson et al., 2003; Hermos et al., 2002; Huizing et al., 2002a), which might result from accumulation of ceroid lipofuscin (an autoﬂuorescent compound) in various tissues (Huizing et al., 2000). The mouse mutants pale ear (HPS1 deﬁcient), light ear

(HPS4 deﬁcient), and double-homozygous pale ear/light ear have identical coat colors and a similar unique hypopigmentation of ears, tail, and feet (Lane and Green, 1967; Li et al., 2004; Suzuki et al., 2002), strongly suggesting a physical and/or functional interaction between HPS1 and HPS4. In addition, tissues of the light ear mouse (HPS4 deﬁcient) show no HPS1 protein expression (Suzuki et al., 2002). A similar phenomenon is seen in cells deﬁcient in subunits of AP3 (Dell’Angelica et al., 1999; Huizing et al., 2002b) or BLOC1 (Ciciotte et al., 2003; Falcon-Perez et al., 2002; Starcevic and Dell’Angelica, 2004), in which the absence of one subunit leads to the loss of other subunits of the corresponding complex. Even though HPS1 and HPS4 are part of a stable complex, no direct interactions between the two proteins have been shown, indicating that other unidentiﬁed BLOC-3 subunits might exist (Martina et al., 2003; Nazarian et al., 2003). In addition, the possible existence of BLOC-4 and BLOC-5 complexes, sharing some BLOC-3 subunits, has been suggested (Chiang et al., 2003). The biochemical pathway in which BLOC-3 functions remains speculative. The HPS1 and HPS4 proteins contain no recognizable homology or structural motifs that can help to predict BLOC-3 function. In human melanotic cells, the HPS1 protein resides in uncoated vesicles and early-stage melanosomes, and the lack of HPS1 protein expression in these cells results in mislocalization of tyrosinase and Tyrp1 to large membranous structures rather than melanosomes, thereby affecting melanin synthesis (Sarangarajan et al., 2001). Subcellular fractionation studies have revealed that BLOC-3 expression is mostly cytosolic, with only a small fraction associated with membranes (Martina et al., 2003). Light ear and pale ear mice have enlarged kidney lysosomes containing increased amounts of lysosomal hydrolases and decreased secretion of lysosomal hydrolases into the urine (Meisler et al., 1980; Swank et al., 1998), suggesting BLOC3 involvement in lysosomal biogenesis. Pale ear ﬁbroblasts exhibited a reduced perinuclear localization of lysosomal and late endosomal markers compared with control ﬁbroblasts, indicating a role for BLOC-3 in the movement and/or localization of these organelles (Nazarian et al., 2003). Finally, ultrastructural studies of eye melanosomes have shown abnormal macromelanosomes within the choroids of pale ear and light ear mice, presumably a result of abnormal melanosome fusion events (Li et al., 2004). The LYST protein, defective in human CHS and in the beige mouse, might also serve as a chaperone in the trafﬁcking of tyrosinase family members to the premelanosome (Barbosa et al., 1996; Huizing et al., 2001b; Nagle et al., 1996; Perou et al., 1997; Ward et al., 2002). LYST is a large cytoplasmic protein (425 kDa) (Nagle et al., 1996), thought to function as an adaptor that may juxtapose proteins that mediate intracellular membrane fusion reactions (Tchernev et al., 2002). LYST contains a BEACH motif, which is highly conserved in a large family of eukaryotic proteins and is crucial for their functions in vesicle trafﬁcking, membrane dynamics, and receptor signaling (Huizing et al., 2001b; Nagle et al., 1996; Ward et al., 161

CHAPTER 7

2002). LYST also contains a series of hydrophobic helices resembling HEAT and ARM domains (Huizing et al., 2001b; Nagle et al., 1996; Ward et al., 2002). HEAT repeat proteins are involved in vesicle transport, and ARM domains are thought to mediate membrane associations (Huizing et al., 2001b; Peifer et al., 1994; Ward et al., 2002). The carboxylterminus of LYST contains several WD40 repeat motifs, which can function in protein–protein interactions (Huizing et al., 2001b; Neer et al., 1994; Ward et al., 2002). Melanocytes in CHS frequently produce large premelanosomes that can reach 10 times the size of a normal melanosome and express an aberrant trafﬁcking of tyrosinase (Zhao et al., 1994). Griscelli syndrome (GS) is associated with defects in three distinct proteins, myosin 5A (GS1), rab27A (GS2), and melanophilin (GS3), the orthologs of the genes mutated in the dilute, ashen, and leaden mice respectively (Anikster et al., 2002; Griscelli et al., 1978; Menasche et al., 2000, 2003; Pastural et al., 1997; Sanal et al., 2002). Myosin 5A, GTP-rab27A, and melanophilin are the main components of a complex that regulates actin-based melanosome transport in melanocytes (Fukuda et al., 2002; Menasche et al., 2003; Wu et al., 2001), as presented in the following Chapter 8. Melanophilin can bind to actin ﬁlaments through its actinbinding domain and serves as the linker protein between myosin 5A and GTP-rab27A (Fukuda et al., 2002). A defect in any component of this complex results in failure of melanosomes to localize to the dendritic tips of melanocytes, thereby preventing melanosome transfer to adjacent keratinocytes (Boissy, 2003; Fukuda et al., 2002; Wu et al., 2001). Several other protein complexes are beginning to emerge as chaperones in melanosome biogenesis. All eukaryotes are known to utilize a combination of coat-associated proteins, SNAREs, syntaxins, rabs (GTPases of the ras superfamily), cytoskeletal proteins, and membrane lipids for vesicle targeting, transport, and fusion events (Karcher et al., 2002; McMaster, 2001; Pelham, 2001; Seabra et al., 2002; Teng et al., 2001). Apart from rab27A, several other rabs also appear to function in melanosome-speciﬁc trafﬁcking, budding, and fusion events (Seabra et al., 2002; Whyte and Munro, 2002). For example, the chocolate coat color mutant mouse, defective in rab38, has impaired targeting of Tyrp1 (Loftus et al., 2002). Rab7, which is localized to late endosomes, is also thought to trafﬁc tyrosinase and Tyrp1 from the TGN to melanosomes (Gomez et al., 2001; Hirosaki et al., 2002), and the HPS mutant mouse gunmetal is mutated in rabggta, an enzyme that attaches geranylgeranyl groups to rabs (Detter et al., 2000). SNARE complexes, which can be either in a vesicle (vSNARE) or in the vesicle’s fusion target (tSNARE), consist of a syntaxin, a vesicle-associated membrane protein (VAMP), and synaptosome-associated proteins (SNAPs) (Pelham, 2001; Teng et al., 2001). The role of the SNARE complex in melanosome biogenesis is slowly becoming apparent. The pallid mutant mouse protein pallidin, a component of BLOC1, interacts with the tSNARE syntaxin 13 (Huang et al., 162

1999). This complex may function as a premelanosomal tSNARE that assists speciﬁc Tyrp1- and Tyrp2-containing vesicles to fuse with the premelanosome (Falcon-Perez et al., 2002; Marks and Seabra, 2001). Other melanosomeassociated SNAREs include syntaxin 4, SNAP-23, and SNAP25 (Scott and Zhao, 2001). Interestingly, the recently described BLOC-1 component Snapin (Starcevic and Dell’Angelica, 2004) can interact with both SNAP-23 and SNAP-25 (Buxton et al., 2003; Ilardi et al., 1999). The so-called ‘granule group’ of Drosophila melanogaster eye color mutants encodes proteins that function in the delivery of proteins to specialized eye pigment granules (Lloyd et al., 1998). Drosophila homologs of all four subunits of AP3 correspond to granule group mutants (garnet/AP3-d, carmine/AP3-m, orange/AP3-s, ruby/AP3-b) (Kretzschmar et al., 2000; Mullins et al., 2000; Ooi et al., 1997), supporting the concept that ﬂy pigment granule defects are related to melanosomal defects in higher organisms. Other proteins defective in the granule group mutants are VPS41 (light), VPS33 (carnation), and VPS18 (deep orange) (Sevrioukov et al., 1999; Shestopal et al., 1997; Warner et al., 1998). Mutations of any of these proteins in yeast result in the accumulation of prevacuolar multiple vesicular bodies (Peterson and Emr, 2001), as in deep orange/VPS18 (Shestopal et al., 1997) and carnation/VPS33 pigment granules (Sevrioukov et al., 1999). VPS41 interacts with the d subunit of AP3 and is required for the formation of AP3-coated carrier vesicles (Rehling et al., 1999). Despite the impressive progress already made in understanding intracellular vesicle formation, a great deal remains unknown. Mutant animal and human pigmentation models provide an invaluable resource for elucidating the pathways involved in the formation and precise trafﬁcking of melanosomal components. These pathways may have broader applicability to cell types other than the melanocyte.

The Melanosome as a Specialized Microenvironment Within the pigment-producing melanosome, TRPs ultimately form into a high-molecular-weight, multimeric complex that acts as a functional, stable, enzymatic unit (Halaban and Moellmann, 1990; Hearing et al., 1982; Jimenez-Cervantes et al., 1998; Orlow et al., 1994). Because of the different mechanisms responsible for the trafﬁcking of TRPs to the melanosome, as described above, this complex presumably does not form until all TRPs are inserted into the limiting membrane of the melanosome. In addition to the tyrosinase gene family members, many other gene products are putatively part of the melanosome and contribute to its specialized microenvironment, allowing melanin synthesis. The luminal acidity of the melanosome appears to regulate melanization. The catalytic activity of tyrosinase exhibits a lag at neutral pH that is lost under acidic conditions (Devi et al., 1987; Lerner et al., 1949; Tripathi et al., 1988), and acidity

BIOGENESIS OF MELANOSOMES

has been shown to control the rate of melanization and tyrosinase activity (Ancans et al., 2001; Saeki and Oikawa, 1983). Other investigators have demonstrated that tyrosinase activity is optimal at neutral pH and reduced in acidic conditions (Hearing and Ekel, 1976; Saeki and Oikawa, 1985; Townsend et al., 1984). Regardless, ﬂuorometric (Bhatnagar et al., 1993) and histocytochemical (Moellmann et al., 1989, personal communication) methods indicate that melanosomes have an acidic lumen in both melanoma cells and normal human melanocytes. Experimental manipulation of light vs. dark skin-derived melanocytes, using inhibitors of the vacuolar proton pump V-ATPase, cause a dramatic increase in tyrosinase activity in light melanocytes and no change in dark melanocytes, without a concomitant alteration in tyrosinase amount (Fuller et al., 2001). This demonstrates that acidity in the melanocyte regulates the efﬁciency of tyrosinase activity. Translocation of tyrosinase out of the RER is impeded in amelanotic melanoma cells exhibiting aberrant vacuolar ATPasemediated proton transport; the defect was corrected by treatment with V-ATPase inhibitors (Halaban et al., 2002). Sodium–hydrogen exchangers (NHEs) are known to regulate intracellular pH (Counillon and Pouyssegur, 2000; Orlowski and Grinstein, 1997). Melanocytes treated with a chemical inhibitor of NHEs result in a rapid dose-dependent inhibition of tyrosinase activity (Smith et al., 2004). In addition, melanocytes express ﬁve of the seven isoforms of NHE (Sarangarajan et al., 2001; Smith et al., 2004), several of which, along with the vesicular proton pump V-ATPase, colocalize with Tyrp1 (Smith et al., 2004). It remains unknown whether regulation of acidity occurs in the melanosome, directly affecting tyrosinase, or within the RER and/or translocational compartments, inﬂuencing the maturation of tyrosinase and/or its efﬁcient delivery to the melanosome. The gene product of OCA2 is the P protein, i.e. the pinkeye dilute protein (Lee et al., 1994), a limiting membrane protein with 12 transmembrane domains (Gardner et al., 1992; Rinchik et al., 1993). Brilliant (2001) demonstrated that melanocytes mutant for P exhibit a loss of colocalization of acidic organelles and Tyrp1, suggesting that the P protein acidiﬁes melanosomes (Puri et al., 2000). Veriﬁcation of this ﬁnding is pending. The P protein also regulates processing in the RER. Speciﬁcally, melan-p1 cells, null at the p locus, exhibit an elevation in tyrosinase activity and protein with concomitant secretion of the enzyme after intracellular cleavage (Manga et al., 2001; Potterf et al., 1998). In addition, the P protein has been shown to exhibit a prominent ER localization and to contribute to the correct processing of tyrosinase in the ER (Chen et al., 2002). It is noteworthy that the P protein, as well as membrane-associated transport protein (MATP), was not identiﬁed by protein analysis of Stage II melanosomes (Basrur et al., 2003), suggesting that these proteins function upstream of the melanosome and are not localized to the organelle. Therefore, it appears that the transmembrane, pore-like P protein is not a prominent component of melanosomes. Its contribution in regulating the environment of the RER and/or Golgi cisternae to facilitate folding

and/or trafﬁcking of tyrosinase, and thus its requirement for stabilization of the TRP complex (Lamoreux et al., 1995), have yet to be elucidated. The P protein was originally hypothesized to be the tyrosine transporter essential to import this substrate into the melanosome for induction of melanization. However, poor sequence homologies between P and known tyrosine transporters (Rosemblat et al., 1994) and the lack of a deﬁciency in tyrosine transport in P null cells (Gahl et al., 1995) refute this. Regardless, the melanosomal membrane does indeed contain a transport protein with a high afﬁnity for tyrosine (Gahl et al., 1995; Potterf et al., 1996). Melan-A/MART-1 is a melanocyte-speciﬁc protein currently of unknown function. It was originally identiﬁed in melanoma patients who expressed T lymphocytes reactive to this melanocyte-speciﬁc protein (Coulie et al., 1994; Kawakami et al., 1994). Melan-A/MART-1 is a 22- to 24-kDa transmembrane glycoprotein oriented with the carboxy terminus exposed to the cytosol (Rimoldi et al., 2001). Melan-A/ MART-1 does not contain consensus signals for N-glycosylation, but it is post-translationally acylated (De Maziere et al., 2002), a common ﬁnding among proteins involved in signaling and trafﬁcking mechanisms (Dunphy and Linder, 1998; Mumby, 1997). Localization studies have demonstrated that Melan-A/MART-1 is an integral membrane protein of melanosomes but decreases in amount as the melanosome matures (De Maziere et al., 2002; Kawakami et al., 1997). However, most of Melan-A/MART-1 (i.e. 80%) is present in the TGN area associated with trans-Golgi cisternae, 50-nm vesicles, and coated early endosomes (De Maziere et al., 2002). These latter structures also import Silver/Pmel17, which subsequently helps to direct their development toward a melanosome lineage as opposed to a lysosome lineage (Raposo et al., 2001). The predominance of Melan-A/ MART-1 in the coated early endosome suggests that it may also participate in directing the early biogenesis of melanosomes. Recently, MATP has been identiﬁed as the product of the gene responsible for oculocutaneous albinism type 4 (OCA4) (Newton et al., 2001). MATP has 12 predicted membranespanning regions lacking signiﬁcant similarities to transporters of amino acids or monosaccharides but with one loop exhibiting similarities to a sucrose/proton symporter in plants (Newton et al., 2001). Melanocytes cultured from the murine model for OCA4, underwhite (uw), express normal levels of tyrosinase synthesis but exhibit dramatically reduced levels of tyrosinase activity (Lehman et al., 2000). In addition, uw melanocytes secrete into the medium dark vesicles that contain tyrosinase, Tyrp1 and Dct (Costin et al., 2003). Similar to the P protein, proteomic analysis of Stage II melanosomes did not identify MATP as a component of mature melanosomes (Basrur et al., 2003). Therefore, it appears that MATP may regulate the trafﬁcking of melanocyte proteins from the Golgi to the premelanosome via its inﬂuence on the luminal milieu of trafﬁcking components, in a fashion similar to that described above for the P and Melan-A/MART-1 protein. 163

CHAPTER 7

Endoplasmic reticulum Golgi

Tyrosinase

Silver/Pmell7

lti

v

ic u

lar/s o r ti n g

bo

no

ed

Mu es

so me

t Coa

en d

a osome/premel

dy

o Melanos

me –

e II stag

Clathrins -

Adaptins Molecules for recognition, docking and fusion -

Fig. 7.5. Schematic diagram of melanosome biogenesis. To date, the initial deﬁning event in the biogenesis of a melanosome is the incorporation and processing of Silver/Pmel17 in a cytoplasmic organelle, deﬁning it as a premelanosome. This cytoplasmic organelle was originally thought to be derived from the endoplasmic reticulum. However, current observations suggest that it may be derived from a coated, early endosomal component. Vesicles containing the precursor form of Silver/Pmel17 may be derived from the endoplasmic reticulum, cis-Golgi and/or trans-Golgi network. Once Silver/Pmel17 is processed in the premelanosome (i.e. Stage I melanosome), matrix assembly progresses, and the organelle develops into a Stage II melanosome that is capable of receiving tyrosinase-related protein (TRP) cargo. After their biosynthesis is complete, TRPs exit the trans face of the Golgi via clathrin-coated vesicles, presumably with the aid of adaptins. Cargo vesicles containing TRPs travel along cytoskeletal elements, possibly being diverted through an intermediate compartment (i.e. multivesicular/sorting body) en route to the Stage II melanosome. At the target Stage II melanosome, recognition, docking, and fusion molecules facilitate incorporation of the TRPs (see also Plate 7.1, pp. 494–495).

Summary Deﬁning all cellular and molecular events regulating the biogenesis of the melanosome is still a work in progress (Fig. 7.5). Original observations intimated that precursor structures for premelanosomes are derived from the ER. However, current observations suggest that premelanosomes originate from the 164

endosomal network of components. A deﬁning event in the commitment of cellular structures to premelanosomes is the incorporation of Silver/Pmel17 into vesicles and its subsequent assembly into a matrix scaffold that deﬁnes the Stage II melanosomes. Concomitant with the formation of Silver/ Pmel17-positive organelles must be the incorporation of pumps and transporters essential for the specialized environment within the melanosomes. Upon progression to Stage II melanosomes, this organelle becomes receptive to enzymes and regulatory molecules that coordinate the synthesis of melanins within. The tyrosinase gene family of proteins are trafﬁcked from the Golgi apparatus to Stage II melanosomes, where molecular steps regulate recognition, docking, and fusing that facilitate their incorporation into melanosomes. The pathways and mechanisms of transit for each tyrosinase gene family member are currently being deﬁned. Interestingly, it appears that differential pathways as well as alternate routes may exist for each TRP. Eventually, these cargo proteins are incorporated into melanosomes, putatively as a complex, and pigment is synthesized. Ultimately, the pigmented melanosomes must be translocated down the melanocyte dendrites for transfer to keratinocytes or to confer color adaptation. Many specialized molecules exist on the melanosome to facilitate this process, as described in the following chapter.

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The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

8

Melanosome Trafﬁcking and Transfer Glynis A. Scott

Introduction In prior chapters, the elegant and complex process of melanosome biogenesis and the biochemistry and regulation of pigmentation have been described. However, there would be little beneﬁt in creating this amazing organelle if the machinery to transport it to the dendrite tip, and transfer it to a keratinocyte, was lacking. Indeed, adequate pigmentation of the skin is as dependent upon successful transport and transfer of melanosomes as it is on the formation of the organelle itself. It has long been recognized that melanocytes and keratinocytes interact both in culture and in skin in vivo (Fig. 8.1). This interaction occurs through long arborizing dendrites that result in the transfer of melanosomes to keratinocytes. Whole organelle donation from one cell to a heterologous cell is unique to the melanocyte–keratinocyte unit. Effective melanosome transfer requires a conﬂuence of successful steps that starts with the formation and maintenance of the dendrite, transport of the melanized melanosome from the cell body to the periphery of the dendrite, followed by transfer of the organelle to a keratinocyte. A virtual explosion of knowledge has occurred in the area of melanosome trafﬁcking and transport in the last 10 years. In this chapter, I will describe the current understanding of these two processes and will review the data that led to these conclusions.

Melanocyte Dendrite Formation and Transport on Microtubules: the First Step in the Process For melanosome trafﬁcking and transfer to occur, effective dendrite formation is a prerequisite. Recent studies have shown that dendrite formation is dependent upon actin polymerization, which is controlled by the small guanosine triphosphate (GTP)-binding proteins of the Rho family (Busca et al., 1998; Scott, 2002; Scott and Cassidy, 1998; Scott and Leopardi, 2003). The Rho subfamily of GTP-binding proteins orchestrates a variety of cellular shape changes, including lamellipodia formation (Rac), ﬁlopodia formation (Cdc42), and stress ﬁber formation (Rho), through their ability to regulate actin assembly in response to growth factors (Etienne-Manneville and Hall, 2002). Similar to other small GTP-binding proteins, Rho proteins alternate between active (GTP-bound) and inactive (GDP-bound) forms, a process that is controlled by regulatory associated proteins (Fig. 8.2). Several studies have

demonstrated the crucial importance of two members of the Rho family of proteins, Rac and Rho, in melanocyte dendrite formation. Our studies have shown that the introduction of constitutively active Rac mutants into human melanocytes and B16F1 melanoma cells stimulates dendrite formation, whereas the introduction of C3 botulinum exoenzyme, which inactivates Rho, inhibits dendrite formation (Fig. 8.3; Scott and Leopardi, 2003). We also showed that ultraviolet irradiation (UVR) and melanocyte-stimulating hormone (MSH), known to stimulate dendrite formation, activate Rac (Scott and Cassidy, 1998). Busca et al. (1998) reported that Rho inhibition is responsible for dendrite formation in B16F10 cells. They showed that treatment of B16F10 cells with C3 botulinum exoenzyme stimulated dendrite formation and that constitutively active forms of Rho inhibited the ability of cAMPelevating agents to stimulate dendricity. We also demonstrated a link between the cAMP second messenger system, known to stimulate dendrite formation, and Rac activation and Rho inhibition (Scott and Leopardi, 2003). Treatment of melanocytes with agents known to stimulate cAMP, such as forskolin and dibutyryl-cAMP, resulted in the rapid activation of Rac and inhibition of Rho (Scott, 2002; Scott and Leopardi, 2003). Therefore, melanocyte dendrite formation is dependent upon actin polymerization, which is in turn controlled by the activity of Rac and Rho, which are regulated by the cAMP second messenger pathway. Whether other second messenger pathways regulate Rac or Rho activity in melanocytes has not been determined. Although the Rho family of proteins is clearly important for melanocyte dendrite formation, other proteins have been shown to inﬂuence dendrite formation, including the Rab protein Rab27B (Chen et al., 2002). Rab27B shares 72% identity with Rab27A, and the introduction of dominant-negative mutants of Rab27B into melanocytes resulted in a decrease in the length and number of dendrites. Although the mechanisms of Rab27B-dependent dendrite formation were not investigated, it is likely due to the role of Rabs in facilitating vesicle transport containing proteins necessary for dendrite formation and/or maintenance. Myosin Va, an unconventional myosin motor (see below), has also been implicated in dendrite formation in melanocytes (Edgar and Bennett, 1999) because the introduction of antisense oligonucleotides to myosin Va inhibits de novo dendrite formation in murine melanocytes. Historically, it had long been known that melanocyte dendrites were composed of both actin and microtubules (Moellmann and McGuire, 1975; Moellmann et al., 1973). Treatment of melanocytes with cytochalasin D, which disrupts the actin cytoskeleton, inhibits melanosome trafﬁcking. 171

CHAPTER 8

Fig. 8.1. Melanocytes contact multiple keratinocytes in vivo. Hematoxylin and eosin-stained section of skin showing a melanocyte at the basement membrane with multiple arborizing dendrites in contact with keratinocytes (see also Plate 8.1, pp. 494–495).

Basement membrane

1µm

Hormones GDP

GEF

GDP GTP Rho

GTP

Rho

Pi

GAP Downstream effectors

Cdc42

Filopodia

Rac

Rho

Lamellipodia

Stress fibers

Fig. 8.2. Schematic of regulation and function of Rho GTP-binding proteins. Rho GTP-binding proteins alternate between active (GTP) and inactive (GDP) bound forms that are controlled by regulatory factors including guanine nucleotide exchange factors (GEF) and GTPase activating proteins (GAP).

However, whether this was due to speciﬁc effects on melanosome transport proteins or to disruption of the integrity of the dendrite was not determined (McGuire and Moellmann, 1972). More recent studies have deﬁnitively established that melanosomes are transported along the length of the dendrite on microtubules. Direct visualization of melanosome trafﬁcking by video microscopy reveals a saltatory back and forth movement with a pathlength and speed characteristic of microtubule-based transport. The addition of nocodizole, which inhibits microtubule formation, results in the rapid redistribution of melanosomes to the cell body, but leaves the integrity of the dendrite intact (Scott et al., 2002; Wu et al., 1997). In many ways, melanosome transport in mammalian cells parallels that seen in amphibians, in which pigment granules are transported on microtubules (Rogers 172

et al., 1997). Several studies have identiﬁed the molecular motors involved in melanosome transport on microtubules. Co-localization studies and Western blotting have shown that melanosomes co-localize with dynein (a minus end microtubule motor) and kinesin (a plus end microtubule motor) (Byers et al., 2000; Hara et al., 2000; Vancoillie et al., 2000). Treatment of human melanocytes with antisense oligonucleotides to a dynein results in enhanced anterograde movement of melanosomes from the cell body to the periphery, providing evidence that dynein transports melanosomes centripetally (Fig. 8.4; Byers et al., 2000). Similarly, treatment of melanocytes with antisense oligonucleotides to kinesin heavy chain (Hara et al., 2000) results in the rapid translocation of melanosomes from the periphery to the cell body, providing evidence that kinesin is the motor that transports melanosomes centrifugally on microtubules. Dynein heavy chains bind microtubules and adenosine triphosphate (ATP) and are responsible for producing forces that move the motor and cargo along microtubules. The intermediate chains link dynein to dynactin, which is responsible for cargo binding. The light intermediate chains, three forms of which have been identiﬁed, are responsible for targeting dynein to different organelles. Analysis of dynein on melanosomes suggests that melanosomes are associated with a distinct light chain (Vancoillie et al., 2000). Kinesin is unrelated in sequence to the other known class of microtubule motor proteins, the dyneins, and is thought to perform functions in the cell distinct from the dyneins. Conventional kinesin is a dimer (Hirose and Amos, 1999) consisting of two identical 120- to 130-kDa chains, commonly known as heavy chains. When kinesin is puriﬁed from brain homogenates, two light chains (60–70 kDa) are associated with the dimers (Kull, 2000). It is thought that both dynein and kinesin remain on melanosomes and that motor activity is

MELANOSOME TRAFFICKING AND TRANSFER A

B

C

Fig. 8.3. Human melanocytes become dendritic in response to Rac activation and Rho inhibition. Human melanocytes in culture were microinjected with either C3 exoenzyme (B, which inhibits Rho; see also Plate 8.2B, pp. 494–495) or constitutively active Rac mutants (C, V12Rac; see also Plate 8.2C, pp. 494–495). Melanocytes become markedly dendritic in response to Rho inhibition and Rac activation compared with cells microinjected with dye alone (A; see also Plate 8.2A, pp. 494–495).

regulated to modify the direction of melanosome movement (Gross et al., 2002). This would account for the observed rapid ability of melanosomes to change direction.

Three Mouse Mutants Prove Critical for Understanding Melanosome Trafﬁcking As has been the case for melanosome biogenesis, mouse mutants proved to be critical in furthering our understanding of melanosome transport. The dilute, leaden, and ashen mice all share a similar phenotype: pigment dilution with a pale or silver coat color, and the microscopic identiﬁcation of clumped melanosomes within the hair follicles. These three mice turned out to be the living embodiment of speciﬁc genetic defects in three individual proteins that cooperate in the ﬁnal stages of melanosome trafﬁcking (Table 8.1).

The ﬁrst major breakthrough came with the identiﬁcation of the mouse dilute gene as the unconventional myosin, myosin Va (Mercer et al., 1991). Unconventional myosins are actin-based motor proteins and are widely expressed (Hasson and Mooseker, 1995). Myosin Va has a globular head domain containing the ATP- and actin-binding sites, a “neck” domain, which is the site of calmodulin (or light chain) binding, and a tail domain, which interacts with cargo. The dilute mouse shows pigment dilution characterized by irregular clumping of pigment in the hair, as well as opisthotonic seizures and death, and both melanocytes and neural cells have been shown to express myosin Va. The tail domain of myosin V serves as an anchor for a variety of cargoes, including melanosomes and synaptic vesicles in mammals to vacuoles and messenger RNA in yeast (Provance and Mercer, 1999). Clues to the function of myosin Va came through immunoﬂuorescence colocalization 173

CHAPTER 8

studies in which myosin Va was shown to be localized to melanosomes (Provance et al., 1996). Immunoelectron microscopy further conﬁrmed the co-localization of myosin Va and melanosomes (Wu et al., 1997). Melanocyte dendrites were normal in dilute mice, indicating that the primary defect caused by myosin Va mutation was in melanosome distribution, rather than in dendrite formation. Similarly, melanosome biogenesis was also shown to be normal in dilute mice (Provance et al., 1996). Because myosin Va, like other myosins, is an actin-based motor, the question of how myosin Va functioned in the transport of melanosomes became one of intense interest. One possibility was that myosin Va transported melanosomes along actin ﬁlaments that extend the length of the dendrite. The other possibility was that the myosin Va–melanosome unit was “captured” at the actin-rich tip of the dendrite, thus positioning the melanosome for transfer to keratinocytes. By taking advantage of the ability to observe melanosomes in vivo through their high reﬂectivity under Nomarsky optics, Wu and colleagues used melanocytes cultured from dilute and wild-type melanocytes in combination with agents that disrupt microtubules to determine that myosin Va functions to facilitate the “capture” of melanosomes at the actin-rich tip of the dendrite (Wu et al., 1997). Evidence to support this model included the demonstraSense DNA

Antisense DNA

12h A

B

C

D

12h

Fig. 8.4. Antisense DNA complementary to cytoplasmic dynein sequence disperses perinuclear melanosomes. Sense DNA-treated melanocytes show normal perinuclear distribution of melanosomes at 12 h (A) and 24 h (C). In contrast, antisense DNA-treated cells show a perinuclear melanosome-free zone (arrow) with increased peripheral membrane pigment distribution at 12 h (B) and loss of perinuclear melanosomes and marked net anterograde dispersion of melanosomes at 24 h (D). Photograph used with kind permission of Dr R. Byers (Byers et al., 2000).

tion that actin ﬁlaments are concentrated at the dendrite tip, and that their orientation is essentially random. Melanosome movement in melanocytes cultured from dilute mice, viewed in real time, demonstrated bidirectional movement with an average speed of 1.04 mm/s for centrifugal movement and 1.13 mm/s for centripetal movement. In the dilute mice, which lack myosin Va, melanosome transport along the dendrite was similar to that in wild-type mice; however, peripheral accumulation of melanosomes did not occur. Introduction of the myosin Va tail domain into wild-type mice resulted in a dilute phenotype, with redistribution of melanosomes to the cell body suggesting that the tail domain acted in a dominant-negative fashion. Further analysis of the myosin Va gene revealed that alternative splice variants contain different combinations of exons B, D, and F. Further, it was shown that cargo selection depends upon the splice variant expressed in a cell (Huang et al., 1998). Analysis of mutations in dilute mice have shown that both the tail and the head regions of myosin Va may be mutated and that exon F is responsible for binding of the tail region of myosin Va to melanosomes (Au and Huang, 2002).

Rab27A and Melanophilin: the Next Pieces of the Puzzle Subsequent to the discovery that myosin Va was the product of the dilute locus, and that it functions to allow capture of melanosomes at the tip of the dendrite, the defect in the ashen mouse was identiﬁed as the Rab protein Rab27A (Wilson et al., 2000). Similar to dilute mice, dendrites in ashen mice are normal; however, melanosome transfer to follicular keratinocytes is abnormal. Rab proteins are a large and diverse family of small GTP-binding proteins that are critical for targeting membrane fusion of vesicles from different cellular compartments. How Rabs work is still unknown; however, it is known that they recruit a large group of different proteins (“effector molecules”) to membranes when they are activated by GTP binding. Over 60 Rab proteins have been identiﬁed in humans. Ashen mice also have bleeding abnormalities, suggesting that Rab27A directs the transport of both melanosomes and vesicles in platelets. Based on genetic evidence, many Rab27A effector proteins have been identiﬁed that share the conserved N-terminal Rab27A binding domain and show Rab27A binding activity in vitro, which suggests that the Rab27A subfamily regulates exocytotic pathways in a variety of cell types. At this point, it was clear that myosin Va and Rab27A both functioned in directing melanosome transport to the dendrite tip, but whether they worked in parallel yet independent

Table 8.1. Mouse mutants, their corresponding human homolog, and protein affected in melanosome trafﬁcking. Mouse mutant

Human homolog

Phenotype

Protein affected

Dilute mouse Ashen mouse Leaden mouse

Griscelli syndrome I Griscelli syndrome II Griscelli syndrome III

Hypopigmentation and neurologic disorder Hypopigmentation and immunologic abnormality Hypopigmentation

Myosin Va Rab27A Melanophilin

174

MELANOSOME TRAFFICKING AND TRANSFER

pathways or as a complex was unknown. When the genetic defect in the leaden mouse was identiﬁed as melanophilin, the product of the MLPH gene, genetic and biochemical techniques were utilized to show that these three proteins, myosin Va, Rab27A, and melanophilin, formed a complex that functioned to capture melanosomes at the dendrite tip. A series of papers demonstrated that Rab27A associates with the membrane of melanosomes, which then recruits melanophilin, which in turn recruits myosin Va (Fig. 8.5; Barral et al., 2002; Fukuda et al., 2002; Strom et al., 2002; Wu et al., 2002a). Rab27A associates with myosin Va and melanophilin (also known as Slac2-a) through distinct regions of the protein, and melanophilin acts as a linker to bridge the interactions between activated Rab27A and myosin Va. The middle region of melanophilin directly binds the globular tail of myosin Va and adjacent exon F (Fukuda et al., 2002; Provance et al., 2002; Strom et al., 2002; Wu et al., 2002b). Melanophilin binds to Rab27A via its N-terminal Slp homology domain. Melanophilin has also been shown to bind actin, and the absence of the actin binding domain of melanophilin results in accumulation of melanosomes in the cell center, adding further support for the “capture model” of melanosome transport originally proposed by Wu and colleagues (Fukuda et al., 2002; Wu et al., 1997). The ability of melanophilin to bind actin, in addition to its function as a Rab27A and myosin Va linker protein, is postulated to transfer melanosomes from microtubules to actin ﬁlaments and may help melanosomes bind to peripheral actin near the tip of the dendrite (Fukuda et al., 2002). In retinal pigment epithelial cells, a similar tripartite complex of Rab27A, MyRIP, and myosin VIIA has been described (El-Amraoui et al., 2002). Rab27A has also been shown to bind to Gnph (involved in insulin secretion) and several other novel proteins of which the function is still unknown (JFC1/Slp1, Slp2-a, Slp3-a, and Slac2-b) (Strom et al., 2002).

Griscelli Syndrome Griscelli syndrome (GS) is an autosomal recessive genodermatosis characterized by hypopigmentation of the skin and hair, clusters of pigment in the hair shaft, and either a primary neurologic impairment or severe immunodeﬁciency. Griscelli syndrome and three deﬁned variants (I–III) are caused by mutations in genes that regulate vesicle transport and are the human homologs of the dilute, ashen, and leaden phenotypes. While all patients with Griscelli syndrome share pigment dilution, they are differentiated by secondary manifestations including mental retardation, developmental delay, and neurologic abnormalities in GS type I and severe immune abnormalities including T cell and macrophage activation, leading to organ inﬁltration and damage in GS type II. The phenotype of patients with GS type III is restricted to hypopigmentation. Griscelli syndrome type II, the largest group, is the human homolog of the ashen phenotype and is due to mutations in Rab27A. Patients exhibit partial albinism, hemophagocytic syndrome, and immune abnormalities. Griscelli syndrome type I is due to mutations in myosin Va

Myosin Va

Actin Head

Proximal tail

Neck Globular tail NH2

Melanophilin

COOH

Rab27A GTP

Melanosome

Fig. 8.5. Schematic representation of the tripartite protein complex responsible for melanosome transport. GTP-bound Rab27A is associated with the melanosome and binds to melanophilin at the amino-terminal domain of melanophilin. Melanophilin binds the globular tail of myosin Va through the C-terminal domain. The head (motor) domain of myosin Va interacts with actin ﬁlaments.

and manifests with partial albinism and neurologic abnormalities. While these two subtypes are generally accepted, there is some evidence that GS types II and I may both represent defects in Rab27A. A report identiﬁed a Muslim Arab kindred with mutations in Rab27A, but not myosin Va, who exhibited variable neurologic abnormalities as well as hemophagocytic syndrome (Anikster et al., 2002). The authors propose that previously reported patients with defects in myosin Va who exhibited neurologic defects but no immune abnormalities may instead have Elejalde syndrome, a condition characterized by mild hypopigmentation and severe, primary neurologic abnormalities, rather than Griscelli syndrome (Anikster et al., 2002). Griscelli syndrome type III is due to mutations in melanophilin (Anikster et al., 2002; Menasche et al., 2000, 2003a), but may also be seen in patients with deletion mutations in exon F of myosin Va (Menasche et al., 2003b).

Melanosome Transfer to Keratinocytes Having produced a melanosome complete with enzymes necessary for melanin synthesis, and having successfully transported it to the tip of the dendrite, the ﬁnal duty of the melanocyte is to transfer the melanosome to the keratinocyte. It is generally agreed that UVR and the hormone MSH stimulate melanosome transfer, although the proteins and 175

CHAPTER 8

signaling intermediates involved in UVR- and MSH-activated melanosome transfer are still unknown (Virador et al., 2002). Other agents, such as niacinamide, the physiologically active amide of niacin, have been shown to suppress melanosome transfer (Hakozaki et al., 2002). A major hurdle that has severely limited progress in understanding the molecular basis of melanosome transfer has been the lack of a model system. The majority of studies of melanosome transfer to keratinocytes have been based on cocultures of nonhuman cells observed by electron microscopy. Studies utilizing time-lapse video microscopy have been limited by the relatively poor resolution achieved (Cohen and Szabo, 1968; Mottaz and Zelickson, 1967; Wolff, 1973). Other more recent studies have utilized gold particle uptake by keratinocytes, melanin uptake, or transfer of cytoplasmic dyes from melanocytes to keratinocytes to measure transfer (Minwalla et al., 2001; Seiberg et al., 2000; Sharlow et al., 2000). In toto, these prior studies led to observations that suggested phagocytosis of melanocyte dendrites by keratinocytes as the major mode of melanosome transfer, although exocytosis of melanosomes into the extracellular space with uptake by keratinocytes and insertion of melanocyte dendrites and transfer of melanosomes to keratinocytes have also been proposed (Yamamoto and Bhawan, 1994). In this chapter, I will ﬁrst review current information about melanosome transfer at the structural level and then discuss data in which molecular mediators of melanosome transfer have been identiﬁed.

Melanosome Transfer Occurs Along Filopodial Extensions of the Melanocyte Dendrite and Cell Body We used high-resolution movies made from digital images of human melanocyte–keratinocyte cocultures to observe directly melanosome transfer to keratinocytes in vitro (Scott et al., 2002). These movies, along with electron microscopy of cells in vitro and skin in vivo, provided evidence showing that melanosome delivery to keratinocytes occurs along ﬁlopodia (Figs 8.6 and 8.7). Time-lapse digital microscopy showed the presence of long (up to 16 mm) dynamically active ﬁlopodia arising from melanocyte dendrite tips and the melanocyte cell body, many of which contained melanosomes that were easily visualized under Nomarsky optics (Fig. 8.6). Close analysis of the movies showed that melanosomes moved along the ﬁlopodia and were transferred to the keratinocytes (Fig. 8.7). We also observed ﬁlopodia arising from melanocytes in human skin biopsies by electron microscopy. These novel observations contradicted the long-held belief that whole tips of dendrites, relatively large structures, are engulfed by keratinocytes. Another important observation was that melanosome transfer is a relatively uncommon event, occurring perhaps once over the course of 45 min of observation. In many instances, melanosomes within ﬁlopodia were observed to travel bidirectionally along the length of the ﬁlopodia, and transfer was often aborted. The ﬁlopodia were observed to attach and detach from the keratinocyte membrane and, in most instances, melanosome transfer did not occur. In a report by Zhang et al. (2004), atomic 176

fp

fp fp

fp A

fp 2µ

2µ

fp

fp

fp

KM

KM

KM

B

C KM

KM

KM

D 2µ Fig. 8.6. Melanocytes exhibit ﬁlopodia. Melanocytes extend long ﬁlopodia from dendrite tips that transport melanosomes to keratinocytes. (A) Two melanocyte dendrites with prominent ﬁlopodia (fp; arrowhead) are shown in images taken at 8 s. The arrowheads point to thin structures consistent with ﬁlopodia. (B) A melanosome (circle) is present in a ﬁlopodium (fp) and moves towards the keratinocyte membrane (KM; outlined by hatched line). Over the course of 16 s, the melanosome has moved along a ﬁlopodium toward the keratinocyte membrane. (C) The tip of a melanocyte dendrite is shown with multiple connections with a keratinocyte membrane (KM; outlined by hatched line). A melanosome (circle) moves toward the KM over the course of 40 s. (D) Two sequential images captured 5 min after treatment of cells with nocodazole are shown. Melanosomes have redistributed toward the melanocyte cell body, leaving a dendrite that appears empty. Filopodia (arrowhead) were not affected by nocodazole treatment.

force microscopy was used to image human epidermal melanocytes. They also noted ﬁlopodia arising from melanocyte dendrites and cell body and the presence of melanosomes within the ﬁlopodia, further establishing ﬁlopodia as a conduit for melanosome transfer. In neural cells, ﬁlopodia primarily function as a guide to the neural growth cone through receptors on its membrane, which responds to exogenous hormones that stimulate growth cone turning and directionality. It is likely that melanocyte ﬁlopodia also function as pathﬁnding structures and that they express receptors that respond to keratinocytederived factors that guide them to the keratinocyte membrane where they attach. While the receptors used for attachment of ﬁlopodia to the keratinocyte membrane are still unknown, work by several groups (Cerdan et al., 1991; Condaminet et al., 1997; Min-

MELANOSOME TRAFFICKING AND TRANSFER A 1

3

2

4

5

6

KC

MD 10µ

2µ

2µ

B

KC MD

10µ

*

Fig. 8.7. Melanosomes are transported to the keratinocyte along ﬁlopodia. (A) A scanning view of a melanocyte dendrite (MD) contacting a keratinocyte (KC) is shown. The boxed area is shown in detail in sequential images taken every 8 s from movies made from this area. A ﬁlopodium arising from the lateral aspect of the dendrite is either attached or inserted into the keratinocyte membrane. A string of melanosomes (approximately six of them) moves in single ﬁle toward the keratinocyte (2–5). (B) A scanning view of a melanocyte dendrite (MD) adjacent to a keratinocyte (KC) is shown. The hatched line delineates the KC membrane in the upper right-hand corner. The boxed area is shown in detail in images that span 152 s. A string of melanosomes (approximately four of them; arrowhead) is present within a projection arising from the side of the body of the melanocyte dendrite. These projections were frequently observed in melanocytes and were shorter and thicker than ﬁlopodia. Other similar projections are present (asterisks).

walla et al., 2001) has shown that lectins and their glycoconjugate ligands may function as receptor–ligand pairs in keratinocyte–melanocyte interactions. Selective lectin binding sites have been demonstrated on melanocytes in normal skin (Nau and Schaumburg-Lever, 1990), and Minwalla et al. (2001) performed functional studies to show that the addition of selected lectins and neoglycoconjugates inhibited melanosome transfer in melanocyte–keratinocyte cocultures.

Molecular Mediators of Melanosome Transfer Direct examination of melanosome transfer shows that melanosomes are trafﬁcked along ﬁlopodia; however, the mechanism by which the melanosome is ﬁnally donated to the keratinocyte is still unclear. One potential mechanism may be exocytosis of melanosomes from ﬁlopodia tips with uptake by keratinocytes through endocytosis, analogous to synaptic vesicle release and uptake in neural cells. Although we did not directly observe exocytosis in digital movies, it is still possible that exocytosis occurs. Virador and colleagues (2002) examined murine melanocyte–keratinocyte cocultures and concluded that exocytosis of melanosomes occurred following UVR or the addition of MSH through determination of melanin content in culture supernatants. However, whether changes in melanin reﬂected an increased number of melanosomes through exocytosis, increased tyrosinase activ-

ity, or sloughing of dendritic tips was not determined. In support of a role for exocytosis in melanosome transfer is the identiﬁcation of SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptors) and Rab3a on melanosomes. SNARE and Rab family of proteins are involved in vesicle transport and docking (Rab) and membrane fusion (SNAREs). The Rab family of GTP-binding proteins regulates vesicle transport as well as exocytosis (for a review, see Jahn and Sudhof, 1999) and localizes to the surface of organelles involved in vectorial transport. Rab3a has been shown to be critical for exocytosis of synaptic vesicles in neurons, secretion from pancreatic b cells, granule secretion in mast cells, and zymogen secretion from pancreatic acinar cells (Takai et al., 1996). Studies have shown that Rab3a, VAMP2, SNAP-23, SNAP-25, and syntaxin-4 are expressed by melanocytes and that Rab3a and SNARE proteins are present on melanosomes (Araki et al., 2000; Scott and Zhao, 2001). These data suggest that SNARE proteins and members of the Rab protein family may be involved in exocytosis of melanosomes. Interestingly, DNA array analysis of genes upregulated by MSH in murine melanocytes showed that several SNARE proteins, including syntaxin-4 and a synaptobrevin-like protein, are upregulated (Virador et al., 2002). Therefore, the presence of SNARE proteins on melanosomes, in conjunction with reports of exocytosis observed ultrastruc177

CHAPTER 8 Serine protease cleavage site

NRSSKGR

Tethered ligand

NH2

NH2 TRYPSIN

COOH COOH

turally or in video capture movies, raises the possibility that exocytosis is a mechanism of melanosome transfer. Another likely mechanism for melanosome transfer is phagocytosis of melanosome-containing ﬁlopodia tips. This hypothesis has been strengthened by work showing that a receptor expressed by keratinocytes, the proteinase activated receptor-2 (PAR-2) is involved in melanosome transfer and skin pigmentation (Seiberg, 2001; Seiberg et al., 2000; Sharlow et al., 2000). The PAR family are G-coupled transmembrane receptors that are activated by serine proteinases, which cleave the extracellular amino-terminal domain of the PAR and expose a tethered ligand that undergoes a conformational change with subsequent activation of the receptor (Fig. 8.8; for a review, see Macfarlane et al., 2001). In HaCat, PAR-2 is activated by trypsin, mast cell tryptase, factor VIIa, or factor Xa, and by synthetic peptides that mimic the amino-terminal portion of the receptor (Camerer et al., 2000; Santulli et al., 1995; Steinhoff et al., 1999). Like all Gcoupled protein receptors, PARs have seven transmembrane domains and associate with heterotrimeric G proteins. PAR-2 is involved in a broad spectrum of physiological processes, including induction of cytokine and prostaglandin expression, mitogenesis, and inﬂammatory responses (Asokananthan et al., 2002; Mirza et al., 1997). Seiberg et al. (2000) and Sharlow et al. (2000) demonstrated that, in keratinocyte–melanocyte cocultures, PAR-2 is activated by UVR, and activation induces melanosome transfer through increased keratinocytic phagocytosis of melanosomes. Electron micrographs of PAR-2 activated cells demonstrated complex membrane rufﬂing and plasma membrane extensions compared with control cells, and PAR-2 activation induced increased actin polymerization and a-actinin ﬁlament organization beneath the plasma membrane (Sharlow et al., 2000). The importance of the actin cytoskeleton in PAR2-mediated melanosome uptake was shown by experiments in which activation of PAR-2 resulted in activation of Rho, which has been implicated in phagocytosis in other cell types (Qualmann and Mellor, 2003; Scott et al., 2003). Activation of PAR-2 has also been demonstrated to alter pigmentation via modulation of melanosome uptake in vivo. Serine protease inhibitors that interfere with PAR-2 activation, such as soybean 178

Fig. 8.8. Schematic representation of PAR-2 activation. PAR-2 is a seven transmembrane receptor that is activated by serine proteases such as trypsin. These act through cleavage of a speciﬁc sequence at the amino-terminal domain of the PAR, which results in unmasking of a new N-terminus that then folds back onto the receptor and activates it. Activation may also be achieved through the use of synthetic peptides (such as SLIGRL-NH2) that mimic the tethered ligand sequence.

trypsin inhibitor, induced a dose-dependent depigmentation of the skin of dark-skinned Yucatan swine. The inhibitors work by inhibiting melanosome transfer and distribution, which leads to skin lightening in vivo (Seiberg et al., 2000a). Moreover, inhibition of PAR-2 activation prevented UVB-induced pigmentation both in vitro and in vivo (Seiberg et al., 2000a). We showed recently that PAR-2 receptor expression is increased in human skin and in cultured keratinocytes following UVR (Scott et al., 2001). Levels of PAR-2 expression, as well as trypsin, which activates PAR-2, are higher in dark-skinned compared with light-skinned individuals (Seiberg et al., 2003). We also showed that UVR-induced PAR-2 expression is attenuated in people who tan poorly (skin type I) compared with people who tan more easily (skin types II and III). Together, these studies suggest that PAR-2 is a critical receptor involved in keratinocyte uptake of melanosomes. Activation of PAR-2 has also been shown to stimulate the release of prostaglandins from human keratinocytes (Scott et al., 2004). Because prostaglandins stimulate dendricity in human melanocytes (Scott et al., in press), as well as melanin synthesis and potentially other aspects of melanocyte biology (Morelli et al., 1989; Nordlund et al., 1986; Parsad et al., 2002; Sauk et al., 1975; Tomita et al., 1992), PAR-2 may control pigmentation through diverse mechanisms.

Conclusion The ability of melanosomes to be trafﬁcked along dendrites and then transferred to keratinocytes is of critical importance for cutaneous pigmentation. It is clear that melanosomes are moved along microtubules, are captured at the dendrite tips through the molecular complex of myosin Va, rab27A, and melanophilin, and are then transferred to keratinocytes along ﬁlopodia. Of the two processes, trafﬁcking and transfer, much more is known about the former than the latter. Future studies will be directed at deﬁning in more precise detail the adhesion molecules that function to regulate melanocyte dendrite and ﬁlopodia attachment to the keratinocyte membrane, the molecules involved in the regulation of phagocytosis of

MELANOSOME TRAFFICKING AND TRANSFER

melanosomes by keratinocytes, and possibly the proteins involved in exocytosis of melanosomes. Still to be determined is the relationship between signaling intermediates, such as the cAMP/PKA pathway, known to stimulate melanosome transfer, and their role in regulating proteins involved in the process of melanosome trafﬁcking and transfer.

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The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

9

Melanosome Processing in Keratinocytes H. Randolph Byers

Summary 1 Melano-phagosomes are transported along microtubule arrays toward the cell center. 2 The motive force of transport of the melano-phagosomes is the microtubule-based motor cytoplasmic dynein. 3 Melano-phagosome fusion with primary lysosomes near the centrosome gives rise to melano-phagolysosomes. 4 Melano-phagolysosomes occasionally form cisternae where melanosomes and dendrite cytoplasm are digested. 5 Melano-phagolysosomes are sometimes distorted into cisternae and stretched into ﬁngerlike projections until individual or small aggregates of melanosomes are pulled off the distorted membrane by dynactin complex/cytoplasmic dynein step motoring along microtubules. 6 Cytoplasmic dynein/dynactin complex maintains the phagocytosed melanosomes and melano-phagolysosomes into perinuclear aggregates in the basal layer and supranuclear melanin caps in the suprabasal keratinocytes. 7 The perinuclear and supranuclear melanin caps (microparasols) protect the nucleus from UV irradiation-induced DNA damage.

Introduction Skin color and protection of the skin from ultraviolet (UV) radiation is primarily dependent on the amount and distribution of melanin in the epidermis. Melanin is the major component of melanosomes, which are produced by melanocytes and transferred into adjacent basal keratinocytes. The suprabasal keratinocytes move upward through progressive keratinocyte layers of the epidermis where melanocyte dendrites continue to transfer melanosomes to the maturing keratinocytes. Melanin is eventually shed with the dead, corniﬁed keratinocytes from the top layers of the stratum corneum. Melanosome transfer is a complex process whereby melanosomes are released or pinched off from the melanocyte dendrite and phagocytosed by adjacent keratinocytes singly or in aggregates. This chapter on melanosome processing in keratinocytes will exclude the rapid advances in our understanding of melanocyte–keratinocyte melanosome transfer mechanisms as they are presented in another chapter (Chapter

8). Therefore, post-phagocytic processing of the phagocytosed melanosome will be the focus of this chapter. The golden age of electron microscopy led to insights into the cellular and ultrastructural relationship of melanocytes, keratinocytes, transference of melanosomes, and the process whereby melano-phagosomes eventually fuse with other primary lysosomes and mature into melano-phagolysosomes (Cohen and Szabo, 1968; Jimbow et al., 1976; Klaus, 1969; Mottaz and Zelickson, 1967; Wolff et al., 1974). More recent immunologic, biochemical, and molecular techniques have accelerated our understanding of melanosome processing in keratinocytes. The close embryologic relationship of cellular derivatives of the neuroectoderm, including neurons, melanocytes, and keratinocytes, has allowed the cellular and molecular insight gained from one cell type to be translated into the other cell type. For example, our understanding of axoplasmic transport has parallel mechanisms in melanosome transport and melano-phagosome transport in keratinocytes. This chapter is divided into two sections, one on the mechanism of keratinocyte melano-phagolysosome distribution and the other on melano-phagolysosome maturation and degradation. The ﬁrst section reviews how elucidating the molecular motors of pigment transport in lower vertebrate pigment cells (chromatophores; including melanophores and erythrophores) led to a deeper understanding of the mechanisms of melano-phagolysosome distribution in human keratinocytes. The second section reviews how our recent understanding of phagosome processing and maturation into phagolysosomes in all eukaryotes and melanocytes has led to insight into the maturation of melano-phagolysosomes in human keratinocytes.

The Mechanism of Melano-Phagosome and Melano-Phagolysosome Distribution in Keratinocytes Early investigators appreciated that the distribution of melanin in the epidermis and keratinocytes is nonrandom (Fig. 9.1). The basal keratinocytes exhibit more melanophagolysosomes than the spinous, granular, and corniﬁed cell layers. The basal keratinocytes show perinuclear aggregation of melano-phagolysosomes around the nucleus. In the suprabasal keratinocytes of tanned individuals or darkly pigmented individuals, melanin is concentrated into “supranuclear 181

CHAPTER 9

Fig. 9.1. Melano-phagolysosomes are organized into supranuclear “caps” within keratinocytes. Note melanized dendritic melanocytes and adjacent keratinocytes with supranuclear “caps.” Melanin silver-stained (Fontana Masson) section of heavily melanized human epidermis. Bar = 50 mm. (Reprinted by permission of Elsevier Science, Inc., from Byers et al., 2003, copyright 2003 by The Society for Investigative Dermatology.)

caps” (Gates and Zimmermann, 1953). This melanin cap is precisely oriented toward the outer skin surface, enhances the protective effect of the nucleus from UV irradiation, and acts as an internal sunscreen. Investigators suspected that, similar to mechanisms responsible for movement of melanosomes observed in melanocytes, a force-generating factor would be found associated with the keratinocyte melano-phagosome. Electron microscopy provided insight into the mechanism of transport of melanosomes in melanocytes but was less helpful in elucidating the mechanism of the melano-phagolysosome distribution in keratinocyte cytoplasm. Only one ultrastructural study speciﬁcally addressed the “forces responsible for the establishment and maintenance of the supranuclear melanin cap” (Findlay and Liebenberg, 1976). These investigators subjected human epidermis to ultracentrifugation and found the melanin cap was relatively shear prone compared with the surrounding intermediate ﬁlaments or tonoﬁlaments (cytokeratin ﬁlaments). This was somewhat of a paradox, as it suggested perhaps a passive displacement or exclusion from the periphery by the abundant intermediate ﬁlaments (cytokeratin ﬁlaments). Still, how would the cytoplasm become organized if it did not involve force generation? Clearly, there is a centripetal force that collects the peripheral phagocytosed melanosomes and transports them to the centrosomes in the perinuclear region. The mechanism of centripetal melanosome movement following phagocytosis in keratinocytes may be traced to our understanding of centripetal aggregation or inward migration of pigment granules in pigment cells of ﬁsh and amphibians. Electron microscopy identiﬁed microtubules oriented in the direction of pigment granule translocation of ﬁsh melanophores, suggesting a role for microtubules in mediating pigment migration (Bikle et al., 1966). Indeed, the dramatic light microscopic radial orientation of pigment granules 182

in pigment cells on ﬁsh scales drawn so beautifully near the end of 1800s by Ballowitz (1893) correlated perfectly with the elegant radial organization of microtubules seen with the electron microscope (Green, 1968; Wikswo and Novales, 1969) or by immunoﬂuorescent localization in vitro (Schliwa et al., 1978) and in situ (Byers et al., 1980) with the ﬂuorescent microscope. Microtubules were functionally shown to be involved in the mechanism when investigators found that the microtubule inhibitors colchicine or vinblastine inhibited pigment granule migration (Green, 1968; Malawista, 1971; Wikswo and Novales, 1969), and that inhibition correlated with microtubule disappearance (Wikswo and Novales, 1972). These investigators occasionally identiﬁed cytoplasmic bridges connecting the pigment granule with the microtubule. However, these bridges were difﬁcult to visualize because of the ultrathin sectioning by conventional electron microscopy. Wholemount cultured pigment cells prepared for the highvoltage electron microscope provided three-dimensional visualization of microtubule to pigment and pigment to pigment ﬁne cytoplasmic bridges (Byers and Porter, 1977). These ﬁne bridges were called the microtrabeculae (Byers and Porter, 1977; Wolosewick and Porter, 1976). This was simply a descriptive term; the motive force appeared to reside within them and, although their molecular nature was completely unknown, speculation centered on acto-myosin-based systems, microtubule-associated proteins, and other uncharacterized cytoskeletal-associated proteins (Byers and Porter, 1977; Luby and Porter, 1980; Schliwa et al., 1981). The ﬁrst microtubule-associated molecular motor characterized was the force-generating dynein arm that produces shear forces in the beating of cilia and ﬂagella (Brokaw, 1994). The hypothesis was put forth that perhaps dynein and microtubules generated the shear forces necessary for translocation of pigment granules (Fujii and Novales, 1969). This mechanism was supported by elegant observations including statistical morphometric analysis of electron micrographs (Murphy, 1975; Murphy and Tilney, 1974) and ingenious use of inhibitors of dynein-like ATPases on ﬁsh pigment cells (Beckerle and Porter, 1982; Clark and Rosenbaum, 1982). Furthermore, dynein hooklet assays on intact and severed dendrites of ﬁsh pigment cells determined that the polarity of microtubules was maintained or repositioned such that their plus ends are oriented peripherally and their minus ends are positioned centrally (McNiven and Porter, 1986; McNiven et al., 1984). The discovery of cytoplasmic dyneins in all eukaryotes (Holzbaur and Vallee, 1994; Milisav, 1998; Schroer, 1994) and the early characterization of cytoplasmic dynein in ﬁsh melanoma cells (Ogawa et al., 1987) implied a role for this microtubule-based motor in pigment transport. Finally, the self-centering of the centrosome in pigment cells and all interphase cells appears to involve microtubule polymerization and formation of the radial microtubule array by cytoplasmic dynein (Burakov et al., 2003; Rodionov and Borisy, 1997; Vorobjev et al., 2001). Cytoplasmic dynein is a double-headed ATPase mechanochemical motor that steps along the microtubule

MELANOSOME PROCESSING IN KERATINOCYTES

toward the minus end of the microtubule at the perinuclear cell center. It functions as a molecular gear mechanism in response to a load by decreasing step distance and increasing force generation (Mallik et al., 2004). The dynein ATPase “nanomotor” heads are composed of two cytoplasmic dynein heavy chains of about 530 kDa each (Holzbaur and Vallee, 1994; Schroer, 1994). Light intermediate cytoplasmic dynein chains of 55 kDa and intermediate chains of 74 kDa bind the motor portion to dynactin, which attaches to membranebound organelles (Gill et al., 1991; Karki and Holzbaur, 1995; Schafer et al., 1994; Vaughan and Vallee, 1995). Heterogeneity of dynein chain subunits with multiple isoforms of the three different chains is hypothesized to participate in coordinated organelle migration or positioning in different cell types (Brill and Pﬁster, 2000; Criswell and Asai, 1998; King et al., 1998; Koonce et al., 1992; Maeda et al., 1998; Mikami et al., 1993; Nurminsky et al., 1998; Paschal et al., 1992; Pﬁster et al., 1996a, b; Tai et al., 2001; Tanaka et al., 1995; Tynan et al., 2000; Vaisberg et al., 1996; Zhang et al., 1993). Indeed, recent evidence suggests that only one of three forms of dynein light intermediate chains fractionates with melanosomes (Reilein et al., 2003). Immunoﬂuorescent and functional blocking studies also point to the role of cytoplasmic dynein in minus-directed melanosome transport. Cytoplasmic dynein co-localizes to melanosomes that are previously stimulated to aggregate in ﬁsh melanophores (Nilsson et al., 1996). Furthermore, microinjection of anti-dynein antibody results in release of aggregated pigment granules and inhibits minus-end microtubule-directed melanosome transport (Nilsson and Wallin, 1997). Similarly, immunolocalization of cytoplasmic dynein shows concentration in the perinuclear region in melanocytes where most of the pigment is positioned (Byers et al., 2000) and localizes precisely with melanosomes (Vancoillie et al., 2000a). Treatment of pigment cells with antisense directed toward the cytoplasmic heavy chain released many of the perinuclear melanosomes (Byers et al., 2000). These studies indicate a role for cytoplasmic dynein in directing transport of melanosomes toward the perinuclear microtubule-organizing centers. Melanosomes, similar to all organelles, are displaced by at least three families of ATPase molecular motors stepping along actin ﬁlaments or microtubule arrays (Lambert et al., 1999; Smith and Simmons, 2001; Vancoillie et al., 2000a). Melanosome transport within human melanocytes is the result of the forces generated by the balance of cell-centered directed cytoplasmic dynein–dynactin microtubule-based transport (Byers et al., 2000; Vancoillie et al., 2000a, b), the outwarddirected myosin V actin-based transport (Provance et al., 1996; Wu et al., 1998), and the peripherally directed kinesin microtubule-based transport (Hara et al., 2000; Vancoillie et al., 2000c). In contrast, in keratinocytes, phagocytosed melanosomes or melanosomal aggregates are rapidly transported to the perinuclear center and tightly maintained there similar to lysosomes (Fig. 9.2) (Byers et al., 2003). Indeed, cytoplasmic dynein knockout results in lysosome dis-

A

B

C

D

E

Fig. 9.2. Time-lapse analysis of cocultures indicates that transferred melanosomes accumulate as perinuclear melano-phagolysosomes in cultured human keratinocytes. (A) Phase-contrast photomicrograph of human melanocyte/keratinocyte coculture. Heavily melanized dendritic melanocytes and keratinocytes with perinuclear melanophagolysosomes are seen. Bar = 50 mm. (B–E) Time-lapse series with 10-s intervals showing distal fragmentation of melanocyte dendrite (see arrows), phagocytosis, melano-phagosome formation, and perinuclear aggregation of melano-phagolysosomes within keratinocytes. Bar = 20 mm. (Reprinted by permission of Elsevier Science, Inc., from Byers et al., 2003, copyright 2003 by The Society for Investigative Dermatology.)

persion in cultured mouse blastocyst cells (Harada et al., 1998). Organelles undergoing transport are linked to the dynactin complex, which in turn is linked to three intermediate chains of cytoplasmic dynein that associate with the two ATPase heavy chains (Hirokawa, 1998). The 74-kDa intermediate chain (Pﬁster et al., 1996a, b) of cytoplasmic dynein is a central link between the organelle and the motor heads stepping along the microtubule. It binds to the p150Glued subunit of the dynactin complex anchored to membrane-bound organelles (Karki and Holzbaur, 1995, 1999; Steffen et al., 1997; Vaughan and Vallee, 1995). Not surprisingly, the 74kDa chain is also present in human melanocytes where it concentrates or co-localizes with pigment granules (Byers et al., 2000; Vancoillie et al., 2000a). The dynactin complex subunits p150Glued and p50 dynamitin also co-localize to melanosomes (Vancoillie et al., 2000b). Similar to human melanocytes, keratinocytes also express the 74-kDa chain and, 183

CHAPTER 9 Sense DNA

Antisense DNA

24h

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Fig. 9.3. Intermediate chain of cytoplasmic dynein co-localizes with perinuclear melano-phagolysosomes and perinuclear microtubules in cultured human keratinocytes. (A) Differential interference contrast image showing perinuclear aggregation of melano-phagolysosomes and other large vesicular organelles compared with small peripheral cytoplasmic vesicular organelles. (B) Anti-tubulin staining showing radial arrangement of microtubules with convergence into perinuclear microtubule-organizing centers. (C) Distribution of the intermediate chain of cytoplasmic dynein is concentrated in the perinuclear region corresponding to the melano-phagolysosomes and other large vesicular organelles with only scant, scattered, small aggregates of peripheral cytoplasmic staining. (D) Overlap image of microtubules and cytoplasmic dynein. Bar = 10 mm. (Reprinted by permission of Elsevier Science, Inc., from Byers et al., 2003, copyright 2003 by The Society for Investigative Dermatology.)

in support of its functional role in the maintenance of perinuclear melano-phagolysosomes, the intermediate chain co-localizes with the perinuclear distribution of melanophagolysosomes and concentrates at the microtubuleorganizing centers of the radial microtubule framework (Fig. 9.3) (Byers et al., 2003). Cytoplasmic dynein is thus in position as the motive force directing the “inward” pull on melano-phagolysosomes to the keratinocyte cell center. In contrast to melanocytes, the inward force on melanosomes in keratinocytes must override the outward “pull” of kinesin (the plus end-directed motor protein family) and myosin V (the actin-based motor protein). Nonetheless, the presence of incomplete overlap of cytoplasmic dynein and microtubules in the perinuclear region (Byers et al., 2003, p. 169) may indicate a role for other proteins in targeting melano-phagolysosomes to the perinuclear aggregation. For example, cytoplasmic dynein light intermediate chain 1 binds to pericentrin, a molecule associated with centrosomes 184

Fig. 9.4. Antisense DNA complementary to cytoplasmic dynein sequence disperses perinuclear melano-phagolysosomes in keratinocytes at 24 and 48 h compared with sense DNA-treated controls. Cocultures of human neonatal melanocytes and keratinocytes (A–C). (A) Sense DNA-treated cells at 24 h show tight perinuclear aggregates of melano-phagolysosomes, whereas antisense DNA treated cells at 24 h (B) reveal perinuclear and numerous dispersed melano-phagolysosomes. (C) Sense DNAtreated cells at 48 h still show tight perinuclear aggregates, whereas antisense DNA-treated cells at 48 h (D) exhibit numerous dispersed melano-phagolysosomes. Bar = 10 mm. (Reprinted by permission of Elsevier Science, Inc., from Byers et al., 2003, copyright 2003 by The Society for Investigative Dermatology.)

at the center of the microtubular array (microtubuleorganizing center). Cytoplasmic dynein also appears to take part in the assembly of pericentrin and g-tubulin-containing microtubule initiation sites (Tynan et al., 2000; Young et al., 2000). Functional evidence for the role of cytoplasmic dynein in retrograde (minus-directed) microtubular melanophagolysosome transport in keratinocytes came from timelapse microscopy of cocultured melanocytes and keratinocytes treated with diluent (control), sense (control), or two different antisense cytoplasmic dynein heavy chain oligonucleotides (Byers et al., 2003). High-resolution time-lapse microscopy allowed visualization of the movement of individual melanosomes or small aggregates of melano-phagolysosomes. Diluent-treated and sense-treated cells showed normal perinuclear aggregation of melanosomes/melano-phagolysosomes, whereas antisense-treated cells exhibited progressive dispersion of the keratinocyte perinuclear aggregates throughout the cytoplasm (Fig. 9.4). Indirect evidence suggests that the microtubule polarities in keratinocytes resemble melanocytes. The antisense experiments on human melanocytes (Byers et al., 2000) infer that the majority of the melanocyte dendrite microtubules are oriented similar to nerve axons (Baas et al., 1988) and ﬁsh melanophores (McNiven and Porter, 1986) with the plus ends directed toward the periphery as opposed to nerve dendrites with mixed polarity (Baas et al., 1988). Similarly, the antisense

MELANOSOME PROCESSING IN KERATINOCYTES

experiments on keratinocytes suggest that the microtubules are oriented with the plus ends in the periphery and the minus ends in the perinuclear centrosome (Byers et al., 2003). Keratinocytes do not show the bidirectional motility of melanosomes that is observed in melanocytes; therefore, it is apparent that the force acting on keratinocyte melanophagolysosomes by the cytoplasmic dynein–dynactin complex overrides any outward-directed forces by myosin V or kinesin motors acting on these organelles. In addition to cytoplasmic dynein’s perinuclear concentration, it is also scattered throughout the cytoplasm, although to a lesser degree, with peripheral punctate staining (Byers et al., 2003). This ﬁnding agrees well with the bidirectional movement of cytoplasmic dynein-associated structures and switching activity of both plus- and minus-directed motors associated with different membrane-bound cargoes (Ma and Chisholm, 2002). Indeed, in order for the melano-phagosome to be pulled from the peripheral cytoplasm, there must be dynactin complexes and cytoplasmic dynein available in the periphery. In fact, cytoplasmic dynein is found at the peripheral membrane in association with adherens junctions containing b-catenin and E-cadherin in epithelia (Ligon et al., 2001). However, the peripheral membrane-associated cytoplasmic dynein that is ready to shuttle cargo toward the cell center is less concentrated than that found in the perinuclear region holding already transported melano-phagolysosomes in place (Byers et al., 2003). Cytoplasmic dynein also has several other functions: the breakdown of the nuclear envelope during prophase (Salina et al., 2002); reorganization of the microtubule array into the mitotic spindle during prophase and metaphase; and possibly chromosomal segregation during anaphase (Banks and Heald, 2001; Barton and Goldstein, 1996; Karki and Holzbaur, 1999; Rusan et al., 2002). Differential expression of members of the dynein superfamily and assignment of speciﬁc isoforms of heavy, intermediate, and/or light intermediate chains may specify cargo trafﬁcking within different cellular compartments and among different cell types. There are at least four cytoplasmic dyneins and 12 members of the axonemal dyneins in the dynein superfamily (Brokaw, 1994; Hirokawa, 1998). Cytoplasmic dynein heavy chain Dyh1 (MAPlc, DHcl, and DHC1a; Koonce et al., 1992; Mikami et al., 1993; Vaisberg et al., 1993; Zhang et al., 1993) is ubiquitous throughout phylogeny from amoeba to vertebrate neurons. In the latter cells, it is responsible for microtubuledependent fast axonal retrograde (minus end-directed) organelle transport (Muresan et al., 1996; Schroer et al., 1989). At least two cytoplasmic dynein family members are present in brain tissue (Tanaka et al., 1995); three have been identiﬁed in ﬁbroblasts and HeLa cells (Vaisberg et al., 1996) and possibly four in rat testis (Criswell and Asai, 1998). Isoform speciﬁcity to cell type or cellular compartment may be involved in different organelle transport, secretion, mitotic spindle organization, chromosome or nuclear migration (Holzbaur and Vallee, 1994; Schroer, 1994; Tanaka et al., 1995; Vaisberg et al., 1996). The cytoplasmic dynein antisense oligonucleotides that induced dispersion of melanosomes from the cell center in melanocytes and

keratinocytes were made to a short Dyh1 sequence cloned from melanocytes (Byers et al., 2000). The lack of complete peripheral dispersion of melano-phagolysosomes in keratinocytes with Dyh1 antisense studies (Byers et al., 2003) suggests functional redundancy of cytoplasmic dynein heavy chain isoforms or functioning of other minus-directed microtubule motors including kinesin-related motors (Case et al., 1997; Henningsen and Schliwa, 1997; Hirokawa, 1998; Matuliene et al., 1999; Noda et al., 2001; Sablin et al., 1998; Smirnova et al., 1998; Wade and Kozielski, 2000). The human epidermis is a polarized epithelium. In tanned or pigmented skin, the supranuclear melanin cap is another feature of polarization. This concentrated aggregate of melano-phagolysosomes over the nucleus of the keratinocyte acts as an internal sunscreen against UV-induced nuclear DNA damage. To emphasize the functional–structural photoprotective effect of aggregation of pigment on a subcellular radial microtubular array over the nucleus, the “microparasol” has been coined (Byers et al., 2003; Scott, 2003). Animal models have demonstrated that the “protective caps” also reside on the side of the nucleus closest to the skin surface and, similar to the tanning response, are increased with aliphatic and alicyclic diols (Brown et al., 1998). The melanin caps of keratinocytes are also seen in vitro with keratinocytes of cultured skin equivalents (Archambault et al., 1995) and increase following UV irradiation or isobutyl methylxanthine (IBMX) treatment (Gibbs et al., 2000). Interestingly, increased supranuclear caps also occur in hyperpigmentation of fullthickness skin grafts (Li et al., 1997), suggesting that UV exposure is not necessary for their formation. In situ immunoﬂuorescent studies also support a role for cytoplasmic dynein as the force-generating motor involved in the formation of the melanin cap. Frozen sections of human skin from a moderately pigmented individual incubated with monoclonal antibodies against the cytoplasmic dynein intermediate chain reveal that cytoplasmic dynein co-localizes with the supranuclear melanin cap (Fig. 9.5) (Byers et al., 2003). Thus, melanin is not randomly distributed throughout the keratinocyte cytoplasm; cytoplasmic dynein forms the supranuclear melanin cap or microparasol to protect the nucleus from UV-induced DNA damage. Earlier studies calculating total melanin content in the skin and testing its protective effect using solutions of melanin are analogous to shredding all the parasols at the beach and spreading them like confetti over the sunbathers and the beach, only to ﬁnd that areas on the bather’s skin would burn, similar to nuclear mutational hits in a basal layer nucleus if the microparasol melano-phagolysosomes were dispersed. The most vulnerable germinative layer, the stratum basale, has the greatest number of melano-phagolysosomes surrounding the nucleus. It may have more pigment surrounding the nucleus as a result of the polarized nature of cell division of daughter cells (one remains on the basal layer and the other moves into the suprabasal layer), and asymmetric segregation of pigment granules may reﬂect a functional anisometry in the melanocyte and keratinocyte cytoskeletal organization. It may also relate to the 185

CHAPTER 9

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Fig. 9.5. Intermediate chain of cytoplasmic dynein in keratinocytes is concentrated in a supranuclear pattern in human epidermis. (A) Phase-contrast image of frozen section of human epidermis showing focal supranuclear aggregates or “caps” of melano-phagolysosomes (arrows). (B) Confocal ﬂuorescence image showing focal increased cytoplasmic staining of the intermediate chain of cytoplasmic dynein in keratinocytes. (C) Double ﬂuorescent confocal image with nuclear staining shows increased supranuclear localization of intermediate chain of labeled cytoplasmic dynein over labeled nuclei (arrows) corresponding to “caps” in (A). (D) Co-localization of melano-phagolysosomes and cytoplasmic dynein intermediate chain by computer-generated pseudo-bright overlay. Bar = 50 mm. (Reprinted by permission of Elsevier Science, Inc., from Byers et al., 2003, copyright 2003 by The Society for Investigative Dermatology.)

temporal sequence of phagocytosis, melano-phagosome transport, processing, and elongation of melano-phagolysosomes into cisterns along the sides of the nuclei prior to maturation of the microparasol. Morphologic and molecular evidence for this is discussed in part below.

Maturation and Degradation of MelanoPhagolysosomes in Keratinocytes Electron microscopic and time-lapse photomicrography studies provided early insights into the processing of melanophagosomes into melano-phagolysosomes. Melanosome size, uptake, and retention as aggregates or complexes were studied as well as the digestion of melanosomes, membranes, and residual melanocyte cytoplasm during maturation into melano-phagolysosomes. Individuals with minimal to lightly pigmented epidermis (phototypes I or II) exhibit small melanosomes packaged within membrane-bound melanosome complexes in the keratinocyte cytoplasm. This contrasts with 186

the single large melanosomes packaged singly in the keratinocyte cytoplasm in individuals with moderately to heavily pigmented epidermis (phototype III or IV) (Hori et al., 1968; Konrad and Wolff, 1973; Szabo et al., 1969). This difference is due to the size of melanosomes produced in the melanocyte; smaller melanosomes are found in phototype I or II and larger melanosomes are seen in phototype III or IV individuals. The size difference inﬂuences the type of transfer and phagocytosis by the keratinocyte (Toda et al., 1972; Wolff et al., 1974; Wolff and Konrad, 1971, 1972). However, recent work indicates a more complex size gradient of melanosomes found in the spectrum of global skin coloration (Thong et al., 2003). In addition, mix and matched co-culture experiments of melanocytes and keratinocytes from dark and light skin have shown that the keratinocytes determine the type of melanosome packaging as either predominantly single melanosomes or predominantly membrane-bound clusters of melanosomes (Minwalla et al., 2001). Nonetheless, depending on the mechanism of transfer and phagocytosis (discussed in Chapter 8), melanosomes are identiﬁed within keratinocytes as: (1) membrane-bound melanosomes as complexes within a pinched-off portion of a membrane-bound melanocyte dendrite and enfolded by keratinocyte membrane; (2) melanosomes enveloped by melanosomal membrane, melanocyte membrane, and keratinocyte membrane to form a trilaminate organelle; and (3) single membrane-bound melanosomes as coated vesicles (Okazaki et al., 1976; Yamamoto and Bhawan, 1994). The general maturation sequence that follows is that keratinocyte primary lysosomes fuse with the melanophagosomes, and the dendrite and/or melanosomes begin to undergo lysosomal digestion. The presence of acid phosphatase in these secondary lysosomes (melano-phagolysosomes) in the keratinocyte supports this pathway (Hori et al., 1968). Nonetheless, melanosomes entering the keratinocyte appear to exhibit a heterogeneous pathway of entrance. In addition, it has become apparent that melanosomes themselves are related to lysosomes in the melanocyte (Orlow, 1995; Zhou et al., 1993). To complicate matters further, melanocytes also exhibit phagocytosis, at least in vitro, of previously released melanosomes (Le Poole et al., 1993). It is thus quite probable that at least a subset of transferred melanosomes may exhibit lysosomal-associated proteins prior to fusion with keratinocyte lysosomes. On the other hand, recent evidence indicates that lysosomes and melanosomes are largely segregated during endosome processing in melanocytes (Fujita et al., 2001; Raposo et al., 2001, 2002; Setaluri, 2003). In addition, the Golgi, lysosomes, and melanosomes are centrally aggregated by similar inward-directed, microtubulebased motor mechanisms involving microtubules and cytoplasmic dynein (Allan et al., 2002; Corthesy-Theulaz et al., 1992; Harada et al., 1998; Karki and Holzbaur, 1999; Valetti et al., 1999). Therefore, caution in the interpretation of mutations or targeted knockout experiments of elements of this mechanism may be in order if a general disruption in the membrane trafﬁcking leads to aberrant membrane fusion and

MELANOSOME PROCESSING IN KERATINOCYTES

mixing of various vesicle compartments that are usually kept relatively separate. The molecular complexity of newly formed phagosomes and mature phagolysosomes is highlighted by the ﬁnding that at least 200 polypeptides associate with these organelles (Desjardins et al., 1994). More recent work has identiﬁed many of the proteins involved in endosome/ lysosome fusion mechanisms and vesicle trafﬁcking, including multiple SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) protein families including syntaxins and VAMPs that bind to SNAPs (SNARE receptors) (Abeliovich et al., 1999; Advani et al., 1999; Atlashkin et al., 2003; Burd et al., 1998; Gerst, 2003; Hay and Scheller, 1997; Houle and Marceau, 2003; Mullins and Bonifacino, 2001; Mullock et al., 2000; Ward et al., 2000). These advances were derived from yeast endosome, lysosome fusion to neuronal synaptic vesicle fusion studies and led to evidence of a role for the SNARE complexes and Rab3a in melanosome transfer (Scott and Zhao, 2001). Undoubtedly, investigators will soon elucidate similar SNARE or Rab GTPase-mediated mechanisms of keratinocyte phagocytosis of melanosomes, melano-phagosome maturation, and transport. In fact, the dynein–dynactin bridging to phagosomes is mediated by Rab GTPase; speciﬁcally, active Rab7 on the phagosomal membrane binds with the effector protein RILP (Rab7-interacting lysosomal protein), which links to the dynein–dynactin complex (Harrison et al., 2003). These investigators observed that phagosomes are transported centripetally, and the dynactin–cytoplasmic dynein microtubule-associated motor complex distorts the membrane into phagosomal tubules toward late endocytic compartments. When RILP was truncated, the tubules did not form, and fusion with late endosomes or lysosomes could not occur. These ﬁndings shed light on the cisternal and tubular membrane structures of late melano-phagolysosomes observed in keratinocytes in earlier electron microscopic investigations (Yamamoto and Bhawan, 1994). Mature melano-phagolysosomes derived from enfolded or sac–dendrite complexes appear to demonstrate two different morphologies: (1) the digestion of the dendrite organelles with fragmentation of the complex and dispersion of individual, pairs, or small aggregates of melanosomes (Okazaki et al., 1976); or (2) digestion of the dendrite organelles with formation of cisternal structures within the keratinocyte (Yamamoto and Bhawan, 1994). These latter cisterns contain electrondense amorphous material and fragments of cytoplasmic components or melanosomes or both. The cisterns become elongate with ﬁngerlike projections containing melanosomes in the tips of the cisterns. The cisterns are often seen running parallel to the nuclear membrane below and to the side of the nucleus, and melanosomes appear to pinch off from the distal cistern. Ultimately, an “apical cap” of aggregated melanosomes may be identiﬁed above the nucleus (Yamamoto and Bhawan, 1994). It is highly probable that the cisterns, as distorted phagosomes and similar to Golgi cisterns, are tubulovesicular components that are pulled and formed by associated cytoplasmic motor proteins acting on microtubules

(Harrison et al., 2003; McIlvain et al., 1993). The exact mechanisms of melano-phagolysosomal maturation and intrakeratinocytic melanosomal membrane trafﬁcking need further clariﬁcation; nonetheless, the elucidation of many basic structural and molecular features provides a framework for the testing of additional hypotheses.

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The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

10

The Regulation of Melanin Formation Vincent J. Hearing

Summary 1 This chapter addresses important regulatory controls of melanogenesis that operate at the subcellular level in melanocytes to affect the structural and/or the functional nature of the melanins and melanin granules produced. 2 Pigment-producing cells in lower vertebrates (usually known as melanophores) are regulated by a number of distinct factors, including hormonal agents such as melanocytestimulating hormone (melanotropin), melanin-concentrating hormone, and melatonin; these generally work by affecting the distribution of existing melanin granules within melanophores. 3 Many of these same factors also affect pigmentation in mammalian melanocytes, but function to regulate the quantity or quality of the melanins synthesized; in addition to melanotropin, the sex hormones (estrogen and progesterone) also affect melanocyte function, and yet another hormone termed agouti signal protein also plays an important physiologic role in regulating melanogenesis in the hair follicle. 4 Melanotropin and agouti signal protein have antagonistic roles and possibly antagonistic mechanisms of action in melanocytes; melanotropin stimulates melanogenic enzyme function and elicits increases in the amount of eumelanins produced; in contrast, agouti signal protein reduces melanin production and elicits the synthesis of pheomelanin rather than eumelanin. 5 There are other levels of controls over pigmentation that are regulated at the melanosomal level, including substrate availability, various cytokines and growth factors, intracellular pH, trafﬁcking and sorting pathways, proteasomes, and ultraviolet radiation, all of which can dramatically affect melanin production and/or the maturation and distribution of melanin granules. 6 The quality and quantity of melanins produced by melanocytes have important physiological consequences for melanocyte function, and undoubtedly play important roles in the various functions of the melanins per se, including hair and skin coloration and photoprotection.

Historical Background Visible pigmentation in the skin, hair, and eyes depends on a

wide variety of factors that inﬂuence melanocyte function at various levels. For correct pigmentation, a number of distinct steps in development, proliferation, and differentiation must occur with high ﬁdelity; these include factors that affect melanoblast development and migration in the developing embryo, affect melanocyte survival and proliferation once in place in situ, affect melanocyte function in response to environmental stimuli, and affect melanin granule distribution and subsequent processing by neighboring keratinocytes. There are a plethora of genes that affect these processes at each of the levels noted, either directly or indirectly. Early on, more than 60 distinct genes were identiﬁed in mice that affect pigmentation (Silvers, 1979), but the number has recently increased to more than 120 (Bennett and Lamoreux, 2003); no doubt that number will increase further in the future. At this time, approximately 50% of those genes have been cloned and characterized, and their general, often their speciﬁc, function(s) in affecting melanogenesis in mammals have been identiﬁed. Several of those genes encode proteins that are localized in melanin granules and play active roles in the structure and function of that organelle, either as catalytic entities involved in melanin synthesis or as structural proteins important to the integrity of the melanosome. Many of those genes, perhaps most of them, have been implicated in various inherited pigmentary disorders in humans, as detailed elsewhere in this book. An excellent resource summarizing the current state of pigment genes, mutations, and associated diseases is maintained online (http://www.cbc.umn.edu/ifpcs/micemut.htm). Most of the genes and factors that modulate melanogenesis at the tissue and cellular levels have been discussed in other chapters of this book and will not be discussed in detail here. This chapter focuses on regulatory controls of melanogenesis that operate at the subcellular level, particularly those that affect the structural or functional nature of melanins and melanin granules produced and/or their processing and distribution within melanocytes. The critical requirement for an enzyme activity to produce melanin from the amino acid tyrosine has been known for some time. The role of tyrosinase [EC 1.14.18.1] was initially described in mushrooms (Bourquelot and Bertrand, 1895; Bertrand, 1896) and, several decades later, tyrosinase was shown also to play a regulatory role in the production of melanin in mammalian melanocytes (Bloch, 1916; Bloch and Schaaf, 1925; Lerner et al., 1949; Lerner and Fitzpatrick, 1950). There was initially some controversy over the identity 191

CHAPTER 10

of the substrate(s) that mammalian tyrosinase uses in melanin production, as the speciﬁcity of the mammalian enzyme was highly restricted compared with tyrosinases present in lower animals and plants. For example, the rate of the initial reaction (tyrosine hydroxylase) proceeds very slowly in the absence of a suitable cofactor, and there was some doubt that tyrosine was the appropriate biologic substrate, but it has now been shown conclusively that tyrosine, via a series of complex reactions, gives rise to the biopolymer melanin. This reaction cascade (discussed in more detail in Chapters 14, 15, and 18) is generally referred to as the Raper–Mason scheme (Raper, 1927; Mason, 1959, 1967); it involves a complex series of oxidations and rearrangements, which results in the formation of indole–quinone ring structures that readily polymerize to high-molecular-weight pigmented biopolymers. Owing to the rate-limiting nature of tyrosinase in this pathway and to the fact that other accessory enzymes were only identiﬁed very recently, early studies assumed that the regulation of melanin production per se was accomplished solely via regulation of tyrosinase activity, and that this in turn was essentially regulated by rates of transcription of its encoding gene. This concept, although in part correct, has been modiﬁed in light of more recent studies. In the past decade, a number of other regulatory points in the melanogenic cascade have been described; these involve inhibitors (which modulate tyrosinase activity), other posttyrosinase enzymes (which regulate the nature of the melanins produced), intracellular pH, proteolytic processing, and other intracellular components that modulate the processing and sorting of melanogenic enzymes, including tyrosinase. As the different types of melanins that can be produced vary greatly in their phenotypic characteristics, it is reasonable to assume that these regulatory factors have great importance to the functional properties of melanins produced. It is important to note that the early concept of direct tyrosinase regulation of melanin formation remains essentially correct, as it is certainly true that, in the absence of active tyrosinase, there will be no melanin formation. However, we now know that there are many ways in which tyrosinase activity is regulated in the melanocyte and that it is not solely at the level of gene transcription; there are many instances in which active tyrosinase is present yet no melanin is produced. We also now know that there are many other switchpoints in the melanin biosynthetic pathway that modulate what type of melanin is produced and how much of it is produced. It should be noted from the outset that we are only now starting to unravel the highly complex series of interactions that is involved in the biogenesis and function of melanin granules, as underscored by the complexity of their proteomic analyses (Kushimoto et al., 2001; Basrur et al., 2003). This chapter can only be considered as a starting point for understanding this ﬁeld, which is an incredibly active area of scientiﬁc research in which important new ﬁndings are being published on a regular basis. 192

Current Concepts Overview It has become increasingly apparent that the pigmentation of mammals and lower animals is under diverse and multifaceted controls. As noted above, there are now a number of factors known to affect pigmentation at the tissue, the cellular, and the subcellular levels, and many of these have been noted and discussed in other chapters of this book. Although the balance of this discussion will focus on the regulation of melanogenesis at the subcellular level, the reader must always keep in mind that regulation at the tissue and cellular levels is equally relevant to the control of pigmentation. As examples, factors that affect the distribution, proliferation, and survival of melanocytes in various tissues are of paramount importance (reviewed by Hearing and King, 1993; King et al., 1997; Spritz and Hearing, 1994); a number of hypo- and hyperpigmentary disorders have been described that result from lesions at this level, including piebaldism and vitiligo. Obviously, if environmental stimuli, e.g. ultraviolet radiation or melanocytestimulating hormone (melanotropin), are operating normally in situ, but functional melanocytes are absent, there will be no pigmentation in that tissue. Similarly, important regulatory effects occur at the cellular level, both within and without melanocytes. Some examples of these types of parameters have also been detailed more completely in other chapters of this book, but they include such cellular determinants as melanocyte structure (e.g. dendrite formation), interactions with keratinocytes (both as a source of regulatory factors and as a repository of donated melanosomes), and other such cellular processes that affect the ultimate distribution and processing of melanosomes in the tissues. Finally, there is active regulation of melanogenesis that occurs at the subcellular level; parameters that fall into this group are those that affect the production and processing of melanin and the melanosomes in which melanin is deposited, at the transcriptional, translational, and/or post-translational levels. Some of these factors are produced within the melanocyte per se, while the vast majority of them derive from outside the melanocyte (e.g. ultraviolet radiation, melanotropin, and agouti signal protein); many of them have pleiotropic effects at different levels, stimulating or inhibiting proliferation of melanocytes in concert with their action in increasing or decreasing pigment production by those same melanocytes. Several of those agents are known not only to modulate the amount of melanin synthesized but also to regulate the type of melanin that is produced (at least in mammals); between those two effects there can be astounding differences in visible phenotype. Chapter 15 discusses in detail the various types of melanin that can be produced, at least in mammals, and the effects on their structure and function. In brief, the two major types of melanins produced are termed pheomelanins and eumelanins, which are yellow/red or black/brown respectively (Fig. 10.1). Interestingly, pheomelanins have not yet been identiﬁed in lower vertebrates and

THE REGULATION OF MELANIN FORMATION

Tyrosinase COOH NH2

HO

Tyrosinase COOH

HO

NH2

HO

Tyrosine

DHI

-O

N H

+

N COOH H DOPAchrome

DOPAchrome tautomerase / DCT O

HO

O

HO

N H Indole-5,6-quinone

NH2

NH2

Glutathione HO or DOPAquinone cysteine N2 H

S

COOH CysteinyIDOPA

HO

COOH N H LeucoDOPAchrome

DHICA oxidase / TYRP1 O N COOH H DHICA

COOH

HO

HO

O

HO

O

DOPA

HO

COOH

O

O

N COOH H Indole-5,6-quinonecarboxylic acid

Eumelanin

COOH

HO

NH2

N S

1,4-Benzothiazinylalanine

Pheomelanin

Fig. 10.1. Schematic of the melanin biosynthetic pathway. DHI, 5,6-dihydroxyindole; DHICA, 5,6-dihydroxyindole-2-carboxylic acid; TRP, tyrosinase-related protein; DOPA, 3,4-dihydroxyphenylalanine.

may represent a more recent and mammalian-speciﬁc group of pigments. Melanotropin and agouti signal protein are factors produced extrinsic to melanocytes that play dramatic roles not only in regulating levels of melanin synthesis, but also in switching melanocytes between production of those two types of melanins, resulting in profound effects on visible pigmentation. Finally, a number of basic housekeeping processes involved in the biogenesis of cellular organelles also regulate the formation and maturation of melanosomes, particularly through the processing and sorting of tyrosinase; such parameters include intracellular pH, proteasome function, and trafﬁcking pathways by which proteins are processed and sorted to their target organelles.

Lower Vertebrates and Invertebrates Pigment-producing cells in lower vertebrates and invertebrates are usually termed melanophores. Similar in morphology and function to mammalian melanocytes, these cells typically appear as highly dendritic cells that are primarily found in the dermis (Bagnara, 1966, 1972; Bagnara and Ferris, 1971; Bagnara and Hadley, 1973; Chavin, 1971; Fujii, 1971; Novales and Novales, 1966). Melanophores are usually present at a higher density on the dorsal surface than on the ventral surface, which in part explains the lighter pigmentation of the latter. However, it has been found more recently that there are also regulatory factors in these tissues, tentatively termed melanization-inhibiting factor and melanizationstimulating factor, which may play important roles in reducing

or promoting pigmentation in these dorsal and ventral surfaces (Bagnara and Fernandez, 1993; Frost, 1988; Fukuzawa and Bagnara, 1989; Fukuzawa and Ide, 1988; Johnson et al., 1992; Wang et al., 2001). These factors have also been shown to have comparable effects on mammalian melanocytes (Kreutzfeld et al., 1989) and may thus play a role in determining the pigmentary patterns in higher vertebrates as well; it is of course possible that some of the melanogenic inhibitors identiﬁed (cf. below and Chapter 18) represent the mammalian counterparts of these stimulating or inhibiting factors. An interesting aspect of melanophore pigmentation is the facility rapidly to translocate melanosomes within the melanophore dendrites, which can lead to relatively quick changes in visible pigmentation based primarily on the distribution of melanin particles within the cell rather than the synthesis of de novo melanin (Bagnara and Hadley, 1973; Fernandez and Bagnara, 1991; Fujii, 2000; Fujii and Novales, 1972; Potter and Hadley, 1970); comparable rapid transit mechanisms of melanosomes in higher vertebrates and mammals have not yet been described. The actual processes and control mechanisms involved in the dispersion and aggregation of melanosomes, and concurrent rates of melanogenesis, in melanophores is discussed extensively in Chapters 2, 3, 7, and 8.

Hormonal Control Pigmentation in lower vertebrates is also under active control by hormones, among which the most commonly known are 193

CHAPTER 10

a – Melanotropin b – Melanotropin g – Melanotropin ACTH (residues 4–10)

MCH (salmon) (rat)

Melatonin

Agouti signal protein

Human–1 Mouse–1

Human–61 Mouse–61

Human–121 Mouse–121 Fig. 10.2. Structure of melanotropic hormones and factors. ACTH, adrenocorticotropic hormone; MCH, melanocyte-concentrating hormone.

melanotropin, melanin-concentrating hormone, and melatonin (Fig. 10.2). In fact, the action of melanotropin was initially deﬁned in amphibians, and early studies in this ﬁeld provided interesting insights into its mechanisms of action (McCord and Allen, 1917; Zondik and Krohn, 1932). Surprisingly, however, more is now known about the signaling pathways and subcellular mechanisms involved in melanotropin action in mammals than in lower vertebrates, primarily because of early studies on the effects of melanotropin in man (Lerner, 1961; Lerner and McGuire, 1961; Lerner et al., 1954; McGuire and Lerner, 1963) and the recent cloning of the family of receptors for melanotropin (Desarnaud et al., 1994; Gantz et al., 1993a, b; Mountjoy et al., 1992). The effect of melanotropin on amphibian melanophores results not only from its effects on the distribution of existing melanin and melanosomes, but also in part from the induction of proliferation of melanophores and/or stimulation of their melanogenic activities, as occurs with mammalian melanocytes (this topic is discussed in more detail in Chapter 2). Melanotropin is a relatively short oligopeptide; it contains 13 residues and is derived from a relatively large precursor termed proopiomelanocortin; this proopiomelanocortin precursor is cleaved proteolytically into a number of hormones, including adrenocorticotropic hormone (ACTH), amelanotropin, b-melanotropin, and g-melanotropin. Proopiomelanocortin (and therefore melanotropin) derives principally from the pituitary gland, but recent evidence has suggested that other cells in the skin (e.g. keratinocytes), as well as melanocytes themselves, can produce proopiomel194

anocortin and may thus act in an autocrine fashion (Chakraborty et al., 1995; Farooqui et al., 1993; Ghanem et al., 1992; Pawelek, 1993; Slominski, 1991a; Slominski et al., 1992, 1995; Wintzen and Gilchrest, 1996). In mammals, a family of melanotropin receptor genes is required for the binding of melanotropin to speciﬁc types of cells and subsequent intracellular signaling (cf. below); receptors for melanotropin have been identiﬁed (de Graan et al., 1981; Eberle, 1981; Fujii and Miyashita, 1976; Graminski et al., 1993; Sherbrooke and Hadley, 1988) but have not yet been cloned in lower vertebrates. In addition to its role in regulating pigmentation in lower vertebrates, melanotropin also plays other roles as a potential neural regulatory agent and growth factor, among other possible functions (cf. below). The mechanism(s) whereby melanotropin effects melanosome dispersion within melanophores has not yet been identiﬁed or characterized; nevertheless, ionic concentrations are important to these responses, especially calcium (Hadley, 1987; McNiven and Ward, 1988; Negishi and Obika, 1985; Veerdonk et al., 1979; Visconti et al., 1989; Visconti and Castrucci, 1990), and no doubt this plays a critical role in the intracellular signaling pathway. The hypothalamus also plays a signiﬁcant role in regulating pigmentation in lower vertebrates (described primarily in teleost ﬁshes) via another hormone known as melaninconcentrating hormone. This hormone essentially has the reverse effect on melanophores that is elicited by melanotropin, i.e. it results in pigment granule aggregation rather than dispersal (Baker, 1988, 1993; Castrucci et al., 1988;

THE REGULATION OF MELANIN FORMATION

Enami, 1955; Hogben and Slome, 1931; Kawauchi et al., 1993; Visconti et al., 1989), and thus lightens rather than darkens visible pigmentation of the skin. Melanin-concentrating hormone is also a short oligopeptide (Fig. 10.2) that consists of 17 residues; the two cysteines present are involved in a disulﬁde bond that cyclizes the molecule (Eberle et al., 1986; Nahon et al., 1993; Okamoto et al., 1984; Wilkes et al., 1984). The effects of melanin-concentrating hormone on the aggregation of pigment granules is reversible, and melanosome dispersion will occur if melanin-concentrating hormone is removed from the milieu or if melanotropin is added (Castrucci et al., 1987; Hadley et al., 1987). The gene encoding melanin-concentrating hormone has been cloned in several different species of salmon (Nahon et al., 1991, 1993; Ono et al., 1988; Takayama et al., 1989) as well as in higher species. As found for melanotropin, melanin-concentrating hormone has a number of alternative functions in amphibians and ﬁshes in addition to its role in eliciting color changes (Baker, 1993; Nahon et al., 1993). Melatonin is a cyclic molecule synthesized from serotonin following acetylation and methylation (Lerner et al., 1958, 1959; Quay and Bagnara, 1964); this synthesis is regulated in the pineal gland and is responsive to light (Bagnara and Hadley, 1973). Melatonin also plays a role in melanosome aggregation in melanophores (Cozzi and Rollag, 1992; Kastin et al., 1972; Messenger and Warner, 1977; Novales, 1963; Rollag and Adelman, 1993), and the responses of pigmentation to variations in light via melatonin are discussed in detail in Chapter 2. The action of melatonin on amphibian melanophores is dependent on the expression of highly speciﬁc melatonin receptors (Heward and Hadley, 1975; Sugden, 2004); indirect evidence suggests that melatonin-speciﬁc receptors also function in mammalian responses to melatonin, although the effect is most likely not on pigmentation per se (cf. below and Chapter 2). Melatonin generally acts in opposition to melanotropin, both directly at the level of the melanophore and indirectly via the hypothalamus, and has been shown to inhibit the effects of melanotropin in amphibians (Abe et al., 1969).

Mammalian More is known about extrinsic regulators of mammalian melanogenesis than for the lower vertebrates, probably because of the added interest in this area due to the cosmetic and pharmacologic implications of the research. This section only brieﬂy discusses hormonal controls and other known extrinsic factors that modify melanogenesis, including ultraviolet radiation and other biologic factors that act in autocrine and/or paracrine manners to inﬂuence the biosynthesis of melanins (processes detailed in Chapters 19, 20, and 21).

Hormonal Control As discussed above for lower vertebrates, hormones such as melanotropin, melanin-concentrating hormone, and melatonin also affect mammalian melanocytes, although often in

distinct manners, yet their physiologic role has often not been resolved. Melanin-concentrating hormone has been cloned and sequenced in mammals and has been shown to be expressed in a tissue-speciﬁc manner, both in rats and in humans (Nahon et al., 1992; Takahashi et al., 1993; Thompson and Watson, 1990). Although the proteins encoded by the mammalian melanin-concentrating hormone genes have been demonstrated to be biologically active (Drozdz et al., 1993), there is still some difference of opinion as to their actual physiologic role in regulating mammalian pigmentation, and as yet there is no direct evidence for such a role. The evidence for possible function(s) of melatonin in the regulation of mammalian melanogenesis is also controversial at best, both for epidermal (McElhinney et al., 1994; Slominski et al., 1994; Snell, 1965; Voulot, 1969) and for follicular (Logan and Weatherhead, 1980; Weatherhead and Logan, 1981a) melanocytes; again, the speciﬁc mechanism(s) of melatonin action, both direct and indirect, in mammalian pigmentation are not yet established. Mammalian melanocytes express speciﬁc surface receptors that bind melatonin, and the encoding gene for that receptor has been cloned and mapped (Garratt et al., 1995). That receptor is expressed by mammalian pigment cells (Roberts et al., 2000), and the speciﬁcity of expression of those receptors has been a source of interest in targeting neoplastic melanocytes (i.e. melanoma); a number of studies have examined this approach (El-Domeini and DasGupta, 1973; Gonzalez et al., 1991; Kane et al., 1994; Weatherhead and Blackburn, 1985). The sex hormones estrogen and progesterone have also been shown to interact with melanocytes, although the mechanism whereby the proliferation or differentiation of melanocytes might be inﬂuenced has yet to be resolved. A number of studies have established that melanocytes express receptors for these hormones (Avril et al., 1985; Chaudhuri et al., 1985; Jee et al., 1994; Walker et al., 1987; Zava and Goldhirsch, 1983), but it is not known whether these hormones have their primary effects on the growth of melanocytes (Beattie et al., 1991; McLeod et al., 1994; Schleicher et al., 1987), activation of the melanogenic pathway (Delacretaz, 1966), and/or speciﬁc binding to the melanins directly (Jacobsohn and Jacobsohn, 1992; Jacobsohn et al., 1993; Kalyanaraman et al., 1986, 1989). It is thought that increases in pigmentation often observed during pregnancy are but one phenotypic effect of estrogen production. Melanotropin, and its precursor proopiomelanocortin, are by far the best characterized of the hormones involved in regulating mammalian pigmentation. Although melanotropin in lower vertebrates regulates pigment dispersion in melanophores (as noted above), in mammals the effect of melanotropin is at least at two other levels, i.e. eliciting an increase in the proliferation of melanocytes (Abdel-Malek et al., 1986, 1987, 1995; Abdel-Malek, 2001; Chavin et al., 1963; Geschwind and Huseby, 1972; Halaban et al., 1993; Halaban and Lerner, 1977; Hill et al., 1989a; Pawelek et al., 1975; Scott et al., 2002a, b; Snell, 1967, 1972; Swope et al., 195

CHAPTER 10

1995; Virador et al., 2002) and/or a stimulation in their production of melanins (Abdel-Malek et al., 2001; Aroca et al., 1993; Burchill and Thody, 1986; Burchill et al., 1986, 1988, 1989; Fuller et al., 1987, 1993; Hadley and Levine, 1993; Hoganson et al., 1989; Weatherhead and Logan, 1981b). The origin of melanotropin that modulates melanocyte function is also an area of active interest — originally proopiomelanocortin was thought to derive only from the pituitary gland, but recent evidence has shown that local production and secretion of proopiomelanocortin by epidermal cells in the skin (Bhardwaj and Luger, 1994; Farooqui et al., 1993; Mazurkiewicz et al., 2000; Pawelek, 1993; Slominski et al., 1992, 1993a, 1993b; Slominski and Wortsman, 2000), particularly keratinocytes (Schauer et al., 1994), may facilitate this paracrine inﬂuence. Perhaps even more interestingly, proopiomelanocortin seems to be expressed by melanocytes themselves and thus acts in an autocrine fashion (Farooqui et al., 1993; Slominski, 1991a; Slominski et al., 1995). There are actually three forms of melanotropin (a, b, and g) that are derived by proteolytic cleavage from the proopiomelanocortin precursor, along with other hormones such as ACTH. Binding of melanotropin to the melanocyte depends upon the surface expression of G protein-coupled melanotropin receptors (Ahmed et al., 1992; Ghanem et al., 1988; Hadley et al., 1981; Siegrist et al., 1989; Varga et al., 1974, 1976). A family of melanotropin receptors (termed melanocortin receptors, MC1R, MC2R, etc.) has been identiﬁed and cloned (Chhajlani and Wikberg, 1992; Chhajlani et al., 1993; Cone and Mountjoy, 1993; Desarnaud et al., 1994; Gantz et al., 1993b, c; Griffon et al., 1994; HaskellLuevano et al., 1994; Konda et al., 1994; Magenis et al., 1994; Mountjoy, 1994; Mountjoy et al., 1992; Siegrist et al., 1994), but only one of these, termed MC1R, is speciﬁcally expressed by melanocytes. MC1R binds the a-melanotropin form of melanotropin preferentially, whereas the distribution patterns and binding characteristics of other members of the melanocortin receptor family differ in other tissues. Presumably, these other receptors have different binding afﬁnities to other members of the proopiomelanocortin derivatives, thus accounting for their varied functions in nonmelanocytic tissues such as the brain. Interestingly, although the MC1R (i.e. the melanocyte-speciﬁc receptor) of most mammals responds only to a-melanotropin, the MC1R expressed by human melanocytes responds equally well to ACTH (Abdel-Malek et al., 2000; Cone and Mountjoy, 1993; Hunt et al., 1994a, b; McGuire and Lerner, 1963), thus explaining the phenotypic effects on pigmentation associated with many conditions in which ACTH metabolism is disturbed. These topics are covered in more detail in Chapters 19, 20, and 21. The intracellular mechanism of melanotropin action is quite complex and not yet completely understood. The melanocytespeciﬁc melanotropin receptor is a G-coupled protein which, upon activation by melanotropin, stimulates the protein kinase A pathway (Abdel-Malek et al., 1992; Busca and Ballotti, 2000; Eberle et al., 1985, 1987; Friedmann et al., 1990a; Haskell-Luevano et al., 1994; Hol et al., 1993; Körner 196

and Pawelek, 1977; Lambert et al., 1985; Mountjoy et al., 1992; Pawelek, 1979, 1987; Varga, 1979). The intracellular events that lead from there to activation of the melanogenic pathway are not fully characterized, but one level of response may involve the stimulation of expression of the transcription factor AP-1, which in turn upregulates the transcription factor Mitf (cf. Chapters 5 and 13) and in turn stimulates the expression and function of melanogenic enzymes, including tyrosinase, and other differentiation factors. Mutations of the melanotropin receptor are associated with pheomelanin production in humans (i.e. red hair and fair skin) (Rees, 2000; Valverde et al., 1995) and can elicit dramatic changes in eumelanin or pheomelanin production depending on whether they lead to hypo- or hyperfunction of the melanotropin receptor (Abdel-Malek, 2001; Abdel-Malek et al., 1999, 2000, 2001; Cone and Mountjoy, 1993; Flanagan et al., 2000; Healy et al., 2000; Magenis et al., 1994; McGuire and Lerner, 1963; Scott et al., 2002a, b). A number of studies have established a correlation between MC1R genotype and melanoma risk (Cone, 2000; Flanagan et al., 2000, 2001; Healy et al., 1999; Rees, 2000; Scott et al., 2002b), although whether that is due to the pheomelanin or via other implications of MC1R signaling is not known. There is no doubt that other responses are also involved in melanocortin receptor signaling and, among those, it seems clear that the protein kinase C pathway is activated in some manner during the melanotropin response and that this also plays a role in the complex series of events that typically leads to stimulation of differentiation and proliferation of melanocytes (Buffey et al., 1992; Englaro et al., 1995; Park et al., 1993; Siegrist et al., 1995). The protein kinase C pathway is yet another route to regulation of melanocyte function and can be the mechanism whereby different growth factors, retinoic acid, and even ultraviolet radiation affect melanogenesis (Ando et al., 1990; Arita et al., 1992; Brooks et al., 1991; Carsberg et al., 1994; Gruber et al., 1992, 1995; Halaban et al., 1992; Loefﬂer et al., 1989; Niles and Loewy, 1989; Oka et al., 1995, 1996; Powell and Rosenberg, 1991; Rosenbaum and Niles, 1992). The last major hormone known to regulate mammalian pigmentation is the latest to be cloned and is in many ways perhaps the most intriguing, i.e. the agouti signal protein, the product of the agouti locus (Fig. 10.2). It has been known for some time that, in mice, melanotropin and agouti signal protein interact to regulate the switch of follicular melanocytes to produce eumelanin or pheomelanin (Fig. 10.3) (Barsh, 1995; Duhl et al., 1994a, b; Granholm and Van Amerongen, 1991; Granholm et al., 1990a, b, 1995; Kucera et al., 1996; Miller et al., 1993; Yen et al., 1994). While melanotropin is produced by a number of cell types in the skin and elsewhere (including melanocytes), agouti signal protein is produced by dermal papillae cells (Millar et al., 1995) and is a paracrine factor that modulates pigment production (Bultman et al., 1992; Miller et al., 1993). Agouti signal protein controls whether black/brown eumelanin or yellow/red pheomelanin is produced by follicular melanocytes, although the precise

THE REGULATION OF MELANIN FORMATION

MC1R

Fig. 10.3. Regulation of melanosomal biosynthesis. MSH, melanocyte-stimulating hormone; ASP, agouti signaling protein; DCT, DOPAchrome tautomerase; gp100, pmel17; MART-1, melanoma antigen recognized by T cells-1; MC1R, melanocortin 1 receptor; MITF, microphthalmia-associated transcription factor; OA1, ocular albinism 1; OCA, oculocutaneous albinism; PKA, protein kinase A; TYR, tyrosinase; TYRP1, tyrosine-related protein 1.

OCA2 OCA4

OCA1 OCA3

Adenylate cyclase G protein

PKA activity

MITF

DCT MART1 OA1

mechanism(s) by which this switch is effected is as yet unknown. In wild-type mice, eumelanin is produced by all follicular melanocytes at the beginning of the hair growth cycle (i.e. from 0–4 days). Transient expression of agouti signal protein from 4–6 days of the hair cycle causes melanocytes to produce pheomelanin instead of eumelanin, after which agouti gene expression is turned off and eumelanin is produced again. This pattern of pigment synthesis results in a yellow striped band near the tip of each hair shaft against a black background (Silvers, 1979; Silvers and Russell, 1955). Eumelanin and pheomelanin differ not only in their gross appearance, but also in their chemical composition and physical properties (as detailed in Chapters 15 and 16) and the ultrastructure of melanosomes in which they are synthesized and deposited. Follicular melanocytes of 1- to 2-day-old agouti mice (i.e. in their eumelanic phase) contain ellipsoid and ﬁbrillar eumelanosomes, while follicular melanocytes of 4- to 6-day-old agouti mice contain ovoid and particulate pheomelanosomes. Similar changes in the ultrastructure of pheomelanosomes and eumelanosomes have been conﬁrmed in a number of mouse mutants that produce one or the other type of melanin (Ozeki et al., 1995; Prota et al., 1995). Mutations at the agouti locus can cause the production of all yellow or all black hair, depending on whether the mutation leads to overexpression/hyperfunction or nonexpression/nonfunction of agouti signal protein respectively (Perry et al., 1994; Siracusa, 1994; Vrieling et al., 1994). Consequently, the agouti locus has been recognized as having an important role in regulating the switch between the production of eumelanin or pheomelanin by melanocytes. The biochemical action of agouti signal protein is a bit controversial (Conklin and Bourne, 1993; Jackson, 1993). Several laboratories have shown (Abdel-Malek, 2001; Abdel-Malek et al., 2001; Blanchard et al., 1995; Lu et al., 1994; Rouzaud et al., 2003; Siegrist et al., 1996; Tota et al., 1999; Willard et al., 1995) that agouti signal protein blocks the ability of melanotropin to activate its receptor, which suggests that the effect of the agouti locus on pigment switching is mediated by

reduced signaling through the melanocortin receptor, whereas its extrapigmentary effects may be mediated by reduced signaling through other melanocortin receptors. However, other groups have shown (Hunt and Thody, 1995; Manne et al., 1995; Zemel et al., 1995) that some effects of agouti signal protein might be mediated by an alteration in calcium channels. Therefore, in addition to working as an antagonist of melanotropin function through the melanotropin receptor, agouti signal protein may also function through other as yet undeﬁned receptors to elicit its pleiotropic effects. Whatever the mechanism, the expression and enzyme activity of tyrosinase in vivo has been shown to be reduced during the pheomelanogenic phase of follicular melanocytes in various genotypes of mice, and there was little or no expression or enzyme activity of tyrosinase-related proteins 1 and 2 during pheomelanogenesis (Furumura et al., 1998, 2001; Granholm et al., 1990b; Kobayashi et al., 1995). This pattern of expression is consistent with the fact that tyrosinase is required for both types of pigment synthesis, but expression of the two tyrosinase-related proteins 1 and 2 is required only for eumelanin synthesis (Kameyama et al., 1992, 1995; Sakai et al., 1997). More recently, a number of studies have shown that melanocytes, which normally produce eumelanin exclusively in vivo, begin to produce signiﬁcant amounts of pheomelanin when cultured in vitro (Hunt et al., 1995a, b; Sakai et al., 1997; Sato et al., 1987), presumably as a result of factors present in the medium. Further, pheomelanin production can be stimulated by in vitro treatment with puriﬁed recombinant agouti signal protein (Hunt and Thody, 1995; Sakai et al., 1997). Following treatment with agouti signal protein, cultured melanocytes exhibit physiologic features characteristic of pheomelanogenesis in vivo, thus providing an in vitro model for future characterization of the mechanisms and genes involved in this switch. Studies using such models should be able to resolve which melanogenic proteins are responsible for the eumelanin:pheomelanin switch and may ﬁnally elucidate the mechanism(s) involved in this important regulatory point. 197

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The function of agouti signal protein in human pigmentation remains speculative at this time (Kanetsky et al., 2002; Krude and Gruters, 2000; Krude et al., 1998; Voisey and van Daal, 2002; Voisey et al., 2001). The homology between the human and mouse proteins is striking and, in fact, the murine and human agouti signal protein elicits pleiotropic effects on murine and human melanocytes in culture (Abdel-Malek et al., 2001; Suzuki et al., 1997; Thirumoorthy et al., 2001). It remains an interesting topic for future study to deﬁne more clearly the role of agouti signal protein in human pigmentation, as well as to characterize its mechanism of action.

demonstrated by several groups (Gahl et al., 1995; Jara et al., 1990a, b, 1991; Pankovich and Jimbow, 1991; Potterf et al., 1996; Saga and Shimojo, 1984). Whether this transport system is constitutively expressed or is actively regulated as a mechanism for controlling melanin synthesis is not yet known, but this seems a reasonable possibility. It also seems quite likely that a distinct membrane transporter will be found to be involved in the control of the pheomelanogenic substrate cysteine through the melanosomal membrane (Potterf et al., 1999) and, again, it will prove interesting to see what role that transporter plays in regulating the switch to pheomelanogenesis.

Substrate Availability

Ultraviolet Radiation

l-Tyrosine, and its hydroxylated derivative l-3,4dihydroxyphenylalanine, are two key substrates of tyrosinase. Increasing the concentrations of these substrates would be expected to increase melanogenesis and, in fact, that does happen. However, such substrate activation of melanogenesis may be biphasic, going beyond the simple substrate activation of enzyme, in that tyrosine and 3,4-dihydroxyphenylalanine can stimulate melanogenesis by stimulating tyrosinase synthesis and activity in the cells (Slominski and Paus, 1990; Slominski et al., 1988). Melanocytes may express receptors that bind the substrates tyrosine and/or 3,4-dihydroxyphenylalanine (Howe et al., 1991; Slominski, 1991b; Slominski and Paus, 1994; Slominski and Pruski, 1992); upon such binding, there is a stimulation of the expression of melanotropin receptors by melanocytes that leads to further increases in proliferation and enzyme activity via melanotropin activation (McLane et al., 1987; Pawelek et al., 1988; Slominski, 1989; Slominski and Costantino, 1991; Slominski et al., 1989a, b). Phosphorylated substrates and other derivatives have been used as an approach speciﬁcally to target melanoma cells (Agin et al., 1987; Pawelek and Murray, 1986). In addition to the possible role of melanogenic substrate receptors on melanocyte differentiation and function, other factors that regulate substrate availability have critical importance to melanin formation. Synthesis of tyrosine itself is dependent upon the action of phenylalanine hydroxylase (Breakeﬁeld et al., 1978; Katz et al., 1976; Nagatsu et al., 1978; Schallreuter et al., 1998; Treiman and Tourian, 1973), yet little is known about the regulation of melanogenesis at this level. In addition, there are substrate transporters that function to regulate melanogenesis at the level of the melanosomal membrane (Gahl et al., 1995; Jara et al., 1990a, b, 1991; Pankovich and Jimbow, 1991; Potterf et al., 1996; Saga and Shimojo, 1984; Tietze et al., 1989). Tyrosine, the principle substrate of the melanogenic pathway, is not freely permeable through biologic membranes and, thus, a tyrosine transporter in the melanosomal membrane seems an obvious requirement. Early studies indicated that the permeability of the melanosomal membrane might play an active role in regulating melanogenic activity (VanWoert, 1970, 1972, 1973; VanWoert et al., 1971) and, in fact, the presence of a speciﬁc tyrosine transporter in the melanosome membrane has been

Ultraviolet radiation is a well-known stimulus of melanogenesis in the skin of mammals, particularly humans, and is commonly known as the tanning response (as detailed in Chapter 17). To summarize the effects with respect to modulation of the melanogenic pathway, the effects noted following ultraviolet radiation are comparable to those elicited by melanotropin, i.e. there is an upregulation in melanocyte proliferation and a stimulation of melanin production (Aberdam et al., 1993; Erickson, 1976; Erickson and Montagna, 1975; Friedmann et al., 1990b; Giacometti et al., 1972; Hunter et al., 1970; Iyengar, 1994; Libow et al., 1988; Morpurgo et al., 1980; Pathak, 1963; Rosdahl and Szabo, 1976; Uesugi et al., 1979; Zelickson et al., 1972). In fact, it has been reported that one mechanism by which ultraviolet radiation elicits its effects on melanocytes is via the induction of expression of melanotropin receptors (Birchall et al., 1991; Chakraborty et al., 1995; Oka et al., 1996; Pawelek et al., 1992), and it has often been noted that melanotropin and ultraviolet radiation produce synergistic effects (Halaban et al., 1993; Seechurn and Thody, 1990). Again, although no direct measurements on quantities and types of melanins produced following ultraviolet radiation of melanocytes have been made (Friedmann and Gilchrest, 1987; Johnson et al., 1972), the upregulation of melanogenic proteins, including tyrosinase and the tyrosinase-related proteins (Allan et al., 1995; Eller et al., 1996; Sharma and Ali, 1976; Sharma et al., 1979), would predict a dramatic increase in eumelanin production. This scenario would be consistent with expectations due to the beneﬁcial ultraviolet-photoprotective properties of eumelanins compared with the more phototoxic properties of pheomelanins (Ahene et al., 1995; Barker et al., 1995; Cesarini and Msika, 1995; Chedekel, 1995; Chedekel et al., 1979; Hill, 1995; Hill et al., 1987, 1989b; Hubbard-Smith et al., 1992; Ishikawa et al., 1984; Jimbow et al., 1995; Menon et al., 1985; Schmitz et al., 1995); this topic is discussed more thoroughly in Chapter 17. Ultraviolet radiation of course can induce different types of photodamage in melanocytes, as well as in other types of cells, which leads to phototoxicity and ultimately to malignant transformation (Donawho et al., 1994; Fenske and Koo, 1994; Kripke, 1974; Nordlund et al., 1981; Setlow et al., 1993; Szabo et al., 1982; Tadokoro et al., 2003; Wolf et al., 1994).

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Other Factors Regulating Pigmentation A number of growth factors are important to melanocyte survival and proliferation; however, as many of these work by stimulating the protein kinase A and/or protein kinase C intracellular signaling pathways, it is not unexpected that they also often have “indirect” effects on rates of melanin production within melanocytes, typically via increased synthesis and activity of tyrosinase (Buffey et al., 1993; Eisinger et al., 1985; Gordon et al., 1989; Halaban et al., 1983). Many of these factors are expressed by neighboring keratinocytes, including basic ﬁbroblast growth factor and endothelin-1 (Ando et al., 1995; Imayama et al., 1994; Imokawa et al., 1995; Nakazawa et al., 1995; Shishido et al., 2001), and their regulation is in turn affected by other environmental stimulants, including ultraviolet radiation (detailed in Chapters 20 and 21). Retinoic acid has been known for some time to upregulate the differentiation (i.e. melanogenesis) and proliferation of mammalian melanocytes (Boskovic et al., 2002; Dupin and LeDouarin, 1995; Edward et al., 1988; Hosoi et al., 1985; Huang et al., 2002; Lotan, 1979; Lotan et al., 1981; Niles and Loewy, 1989; Orlow et al., 1990; Talwar et al., 1993). As with ultraviolet stimulation, this effect seems to be mediated through an increase in expression of melanocortin receptors (Chakraborty et al., 1990, 1991; Gruber et al., 1995; Ho et al., 1992; Niles and Loewy, 1991; Rosenbaum and Niles, 1992). Thus, one might reasonably expect a similar stimulatory pattern of expression as seen with melanotropin itself, i.e. an increase in tyrosinase expression and activity (accompanied by moderate changes in the catalytic functions of tyrosinaserelated proteins 1 and 2) and thus a stimulation of eumelanin production. In addition to factors that upregulate pigment production in melanocytes, there are several physiologically important ones that downregulate melanogenesis. Agouti signal protein is one such notable factor that is produced extrinsic to melanocytes and is discussed above. It is known that such inhibitors play important roles in regulating ventral/dorsal pigmentation patterns in amphibians, and the relationship of agouti signal protein to those factors, as well as the role that differential expression of agouti signal protein plays in regulating mammalian pigmentation patterns, is another interesting question that remains to be examined. The role of melanogenic inhibitors is covered in detail in Chapter 18. Yet another physiologically important inhibitory factor is an intrinsic melanin inhibitor found within murine and human melanocytes, the level of which seems to play an active role in regulating melanin production by existing melanogenic enzymes (Hearing et al., 1995; Kameyama and Hearing, 1990; Kameyama et al., 1989, 1990, 1992, 1995). While tyrosinase is a critical requirement for melanin production, the presence of tyrosinase within melanocytes is no guarantee of melanin synthesis (Aroca et al., 1993; Eller et al., 1990; Hearing and Jiménez, 1989; Jara et al., 1988; Jiménez et al., 1988; Kameyama et al., 1989; Naeyaert et al., 1991), and many amelanotic, but tyrosinase-positive cell lines have been described both in vivo and in vitro. As discussed in Chapters

11 and 12, there are a number of tyrosinase-positive forms of oculocutaneous albinism in which normal levels of active tyrosinase are present, yet little or no melanin is produced. Full chemical characterization of these endogenous inhibitors has remained elusive, and future work will be necessary not only to identify these, but to elucidate their role in the metabolic regulation of melanin production in melanocytes.

Intracellular Trafﬁcking and Melanosome Biogenesis The known melanosomal proteins are all involved in melanogenesis as catalytic and/or structural components of that organelle, and are currently composed of tyrosinase, TYRP1, DCT, MART 1, and gp100 (Kobayashi et al., 1994; Kwon et al., 1991). Although the processing and sorting of these proteins is not completely understood, they are known to be synthesized and translocated into the endoplasmic reticulum and eventually into the Golgi, where their post-translational processing and glycosylation takes place (Francis et al., 2003; Olivares et al., 2003). Following that, they seem to take distinct routes to trafﬁc to melanosomes, the majority of melanosomal proteins predominantly going through the endosomal system to Stage II melanosomes, although that distribution can be disrupted in amelanotic cells. Despite their high structural similarity and conserved primary sequences, the three tyrosinase-related proteins seem to use distinct routes to move from the trans-Golgi network to early melanosomes: tyrosinase uses the AP3 system, TYRP1 uses the AP1 system, and DCT uses yet another unknown sorting vesicle system (Huizing et al., 2002; Negroiu et al., 2003; Spritz et al., 2003). The sorting system for MART 1 is not yet known, but its distribution patterns are highly similar to tyrosinase. gp100 is perhaps the most uniquely processed of the melanosomal proteins; after processing through the endoplasmic reticulum and Golgi, it is normally delivered to Stage I melanosomes without going through the endosomal system (Basrur et al., 2003; Kushimoto et al., 2001; Yasumoto et al., 2004). Following the maturation of Stage I to Stage II melanosomes, which is coincident with the cleavage and refolding of gp100, the enzymatic components are delivered and the synthesis of melanin usually ensues (Kushimoto et al., 2003). Interestingly, the subcellular distribution of melanosomal proteins in amelanotic melanocytic cells that produce normal levels of melanosomal proteins (presumably with intact functional activities) and thus should be pigmented is quite distinct. In pigmented cells, the majority of tyrosinase is correctly processed through the endoplasmic reticulum, Golgi, and endosomes and is delivered to Stage II melanosomes (Kushimoto et al., 2001; Watabe et al., 2004). In contrast, the majority of tyrosinase in amelanotic cells is retained in the endoplasmic reticulum, although a minor amount of tyrosinase can be glycosylated correctly and found in the Golgi, endosomes, and early melanosome fraction (primarily in Stage I melanosomes). Melanosomes are lysosome-related organelles, but their exact biogenesis is still poorly deﬁned (Marks and Seabra, 2001; Raposo et al., 2001). For instance, 199

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melanosomes and lysosomes contain many of the same structural proteins [e.g. lysosome-associated membrane proteins (LAMP), acidic hydrolases, vacuolar-type proton pumps] (Ancans and Thody, 2000; Bhatnagar and Ramaiah, 1998; Diment et al., 1995), and both are affected in several genetic disorders, such as the Chediak–Higashi and Hermansky– Pudlak syndromes (Boissy et al., 1998; Zhao et al., 1994). The correct processing and sorting of tyrosinase to melanosomes is the key to pigmentation and a number of factors regulate it. In addition to pH (as discussed below), other chaperonelike proteins are involved (e.g. TYRP1), as are vesicular components such as the P protein and MATP, and disruptions in the functions of these proteins are associated with oculocutaneous albinism types III, II, and IV respectively (Costin et al., 2003; Toyofuku et al., 2001a, b, 2002) (detailed in Chapters 11 and 12).

Proteasome Function and the Regulation of Tyrosinase Activity In contrast to the distribution and stability patterns of tyrosinase in pigmented cells, the majority of tyrosinase in amelanotic cells is not correctly glycosylated, is trapped in the endoplasmic reticulum, and is quickly degraded by proteasomes. This mechanism is also involved with the hypopigmentation seen in various forms of inherited pigmentary disorders such as oculocutaneous albinism, as noted above. Thus, tyrosinase, which is an extremely stable protein with a half-life of 16–24 h in pigmented melanocytes, can have an extremely short half-life in amelanotic cells, and that stability can be markedly enhanced by treatment with proteasome inhibitors, showing that proteasomes are involved in the degradation of tyrosinase. It remains an interesting hypothesis that proteasomes (and the degradation of tyrosinase) may actually be a physiologically important point of regulating tyrosinase activity and pigmentation under normal circumstances. A recent series of studies has found that free fatty acids can dramatically stimulate or inhibit pigmentation (Ando et al., 1999). Analysis of the mechanism involved in those effects has shown that they speciﬁcally affect tyrosinase and, in fact, they reﬂect the effects of those free fatty acids on proteasome function (Ando et al., 2004; Kageyama et al., 2004). At least one pigment-related gene has been associated with a ubiquitin ligase function in regulating pigmentation (He et al., 2003) and, thus, the role of proteasomes in modulating melanogenesis, perhaps providing rapid responses to environmental stimuli that require no changes in gene expression, is an exciting avenue for future study.

has been some controversy regarding the optimal pH for tyrosinase activity. Melanosomes, like other lysosomal organelles, can be quite acidic, and it has been assumed that this low melanosomal pH facilitates melanogenesis (Devi et al., 1987). It has been suggested that TYR may be activated at an acidic pH and that the enzyme is inactive at neutral pH (Tripathi et al., 1988). However, several other groups have maintained that mammalian TYR has an optimal enzymatic activity near neutral pH and that its activity is gradually lost with decreasing pH (Hearing and Ekel, 1976; Saeki and Oikawa, 1985; Townsend et al., 1984). However, the pH at which melanin is produced and the pH at which the vesicular trafﬁcking system works may function quite independently. TYR levels in amelanotic cells can be dramatically increased following treatment with protonophore or proton pump inhibitors (Fuller et al., 2001; Halaban et al., 2002; Manga and Orlow, 2001; Watabe et al., 2004). Surprisingly, the retention of TYR in the endoplasmic reticulum in amelanotic cells is corrected by these agents (i.e. by increasing intracellular pH), which results in the enhanced transport of tyrosinase from the endoplasmic reticulum to the Golgi. Thus, the abnormal acidiﬁcation of intracellular organelles also plays an important role in the pathogenesis of hypopigmentation in amelanotic cells, and may even play an important role in regulating the constitutive level of pigmentation in the skin. Although tyrosinase is dramatically redistributed to Stage II melanosomes in amelanotic cells following the neutralization of intracellular pH, tyrosinase levels in endosomes are reduced in the presence of proton pump inhibitors but are increased in the presence of protonophores. This suggests that the processing and trafﬁcking of tyrosinase in amelanotic melanocytes is related to the dysfunction of vATPases. vATPase was recently identiﬁed as a melanosomal protein (Basrur et al., 2003) and, thus, it may play an important role in regulating pigmentation. Transport of tyrosinase from the endoplasmic reticulum and its subsequent processing depends on the neutralization of pH in the Golgi, although the translocation of tyrosinase to endosomes then requires the activation of vATPase. Neutralization of the pH within early melanosomes results in the accumulation of tyrosinase in those organelles. Therefore, the activity of vATPases within intracellular organelles plays an important role in the sorting and function of tyrosinase and in modulating pigmentation; dysfunction of this pH regulatory system may be responsible for the depigmented phenotype and pathogenesis of amelanotic melanocytes.

Perspectives Regulation of Melanin Production via Intracellular pH The catalytic domains of tyrosinase and other enzymes involved in melanogenesis are located within the lumen of the melanosome, and it follows that their activity is likely to be dependent upon the intramelanosomal environment, including the pH (Ancans et al., 2001; Puri et al., 2000). However, there 200

The regulation of pigmentation has important consequences for all the varied functions of melanin, including the social implications of skin pigmentation, the cosmetic variations in skin, hair, and eye color, and of course its photoprotective function. It has become progressively obvious in recent years that there are numerous levels of control over pigmentation, which

THE REGULATION OF MELANIN FORMATION I

III

II

IV

Dct Tyr

Fig. 10.4. Melanosomal architecture. Dct, DOPAchrome tautomerase; DOPA, 3,4-dihydroxyphenylalanine; DHI, 5,6-dihydroxyindole; DHICA, 5,6dihydroxyindole-2-carboxylic acid; Tyr, tyrosinase; Tyrp1, tyrosine-related protein 1.

Tyrosine

are exceedingly complex, and that it will be some time before we can fully appreciate the interactions that are involved. Regulatory stimuli for melanocyte function originate from the environment (e.g. ultraviolet radiation), endocrine sources (e.g. melanotropin from the pituitary), paracrine sources (e.g. growth factors, endothelins, or agouti signal protein from keratinocytes or other neighboring cells), or can be autocrine (e.g. proopiomelanocortin from the melanocyte itself). In addition, the pigment distribution depends upon the initial distribution and survival of melanoblasts from the developing embryo and, of course, ultimately in mammals on the transfer and processing of melanosomes to keratinocytes in the skin or hair. Each of these steps is under complex genetic and molecular controls, and it is no wonder that there is such a variety of visible phenotypes in humans. In addition to studies targeted at understanding the regulation of processes involved in pigmentation, the melanosome (Fig. 10.4) is now also being used as a model for study of organelle biogenesis and function as well as for targeting of malignant melanoma. The complexity of controls regulating pigmentation is an impediment to easy understanding of the entire process, but promises a wealth of potential targets for the eventual manipulation of the system, for either cosmetic or pharmacologic purposes.

Acknowledgments The author wishes to thank numerous investigators in this ﬁeld who have been so cooperative in countless discussions on these topics in recent years. Among them are Drs Zalfa AbdelMalek, Gregory Barsh, Dorothy Bennett, Raymond Boissy, Murray Brilliant, Shosuke Ito, Richard King, Lynn Lamoreux, William Oetting, Seth Orlow, John Pawelek, Giuseppe Prota, Francisco Solano, Richard Spritz, and all former/current members of the Hearing Laboratory — I apologize to many others whose names have been omitted for the sake of brevity.

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DOPAquinone

DOPAchrome + cysteine

Tyrp1

DHICA DHI

DHICA-melanins

DHI-melanins

Pheomelanins

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V. J. Hearing. Inﬂuence of a-melanocyte stimulating hormone and ultraviolet radiation on the transfer of melanosomes to keratinocytes. FASEB J. 16:105–107, 2002. Visconti, M. A., and A. M. Castrucci. Ionic requirements for catecholamine aggregating actions on the teleost Poecilia reticulata melanophores. Pigment Cell Res. 3:132–140, 1990. Visconti, M. A., A. M. Castrucci, M. E. Hadley, and V. J. Hruby. Ionic requirements for melanin concentrating hormone (MCH) actions on teleost Poecilia reticulata melanophores. Pigment Cell Res. 2:213–217, 1989. Voisey, J., and A. van Daal. Agouti: from mouse to man, from skin to fat. Pigment Cell Res. 15:10–18, 2002. Voisey, J., N. F. Box, and A. van Daal. A polymorphism study of the human agouti gene and its association with MC1R. Pigment Cell Res. 14:264–267, 2001. Voulot, C. Effect of melatonin and various indoles on the guinea pig melanocyte. C. R. Acad. Sci. 269:2247–2250, 1969. Vrieling, H., D. M. J. Duhl, S. E. Millar, K. A. Miller, and G. S. Barsh. Differences in dorsal and ventral pigmentation result from regional expression of the mouse agouti gene. Proc. Natl. Acad. Sci. USA 91:5667–5671, 1994. Walker, M. J., C. W. Beattie, M. K. Patel, S. M. Roman, and T. K. DasGupta. Estrogen receptor in malignant melanoma. J. Clin. Oncol. 5:1256–1261, 1987. Wang, S. Q., R. B. Setlow, M. Berwick, D. Polsky, A. A. Marghoob, A. W. Kopf, and R. S. Bart. Ultraviolet A and melanoma: a review. J. Am. Acad. Dermatol. 44:837–846, 2001. Watabe, H., J. C. Valencia, K. Yasumoto, T. Kushimoto, H. Ando, J. Muller, W. D. Vieira, M. Mizoguchi, E. Appella, and V. J. Hearing. Regulation of tyrosinase processing and trafﬁcking by organellar pH and by proteasome activity. J. Biol. Chem. 279:7971–7981, 2004. Weatherhead, B., and M. E. C. Blackburn. Growth and melanization of a hamster melanoma cell line, RPMI 3460, in response to melatonin and cyclic nucleotides. In: Biological, Molecular and Clinical Aspects of Pigmentation, J. T. Bagnara, S. N. Klaus, E. Paul, and M. Schartl (eds). Tokyo: University of Tokyo Press, 1985, pp. 471–476. Weatherhead, B., and A. Logan. Interaction of a-melanocytestimulating hormone, melatonin, cyclic AMP and cyclic GMP in the control of melanogenesis in hair follicle melanocytes in vitro. J. Endocrinol. 90:89–96, 1981a. Wilkes, B. C., V. J. Hruby, W. C. Sherbrooke, A. M. Castrucci, and M. E. Hadley. Synthesis and biological actions of melanin concentrating hormone. Biochem. Biophys. Res. Commun. 122:613–619, 1984. Willard, D. H., W. Bodnar, C. Harris, L. Kiefer, J. S. Nichols, S. Blanchard, C. Hoffman, M. Moyer, W. Burkhart, J. Weiel, M. A. Luther, W. O. Wilkison, and W. J. Rocque. Agouti structure and function: characterization of a potent a-melanocyte stimulating hormone receptor antagonist. Biochemistry 34:12341–12346, 1995. Wintzen, M., and B. A. Gilchrest. Proopiomelanocortin, its derived peptides, and the skin. J. Invest. Dermatol. 106:3–10, 1996. Wolf, P., C. K. Donawho, and M. L. Kripke. Effect of sunscreens on UV radiation-induced enhancement of melanoma growth in mice. J. Natl. Cancer Inst. 86:99–105, 1994. Yasumoto, K., H. Watabe, J. C. Valencia, T. Kushimoto, T. Kobayashi, E. Appella, and V. J. Hearing. 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, 2004. Yen, T. T., A. M. Gill, L. G. Frigeri, G. S. Barsh, and G. L. Wolff. Obesity, diabetes, and neoplasia in yellow Avy/- mice: Ectopic expression of the agouti gene. FASEB J. 8:479–488, 1994.

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The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

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The Tyrosinase Gene Family William S. Oetting and Vijayasaradhi Setaluri

Summary 1 The focus of this chapter is the molecular biology of the members of the tyrosinase gene family. Genes of this family encode enzymes involved in the melanin biosynthetic pathway. Members of this family include tyrosinase (the rate-limiting enzyme in melanin biosynthesis), 5,6-dihydroxyindole-2carboxylic acid (DHICA) oxidase, and DOPAchrome tautomerase (also known as TRP1 and TRP2 respectively). 2 All three members of this gene family exhibit high nucleotide sequence homology. This homology extends throughout the animal kingdom. 3 The protein products of these three genes are also highly homologous and share common structural domains including a transmembrane domain, two putative copper binding sites, and a cysteine-rich domain containing an epidermal growth factor (EGF)-like protein-binding motif. 4 The transcriptional regulation of the tyrosinase family genes is rather complex, involving multiple cis-regulatory elements and transactivating factors. A 5¢ regulatory element, termed M-box, that is common to all three genes, is necessary for their regulation by the microphthalmia transcription factor, MITF. However, melanocyte-speciﬁc expression of tyrosinase genes during development and differentiation may involve both positive and negative regulatory mechanisms.

Historical Background The tyrosinase gene family consists of three genes that code for enzymes active in the melanin biosynthetic pathway, and truncated pseudogenes in some species. Tyrosinase is the most well-known member of this gene family and is responsible for the rate-limiting step in melanin biosynthesis — the hydroxylation of tyrosine to dopaquinone (Cooksey et al., 1997). The ﬁrst report of tyrosinase was in 1895, when tyrosinase activity in mushrooms was described as a heat-sensitive, watersoluble ferment that turns black upon exposure to oxygen (Bourquelot and Bertrand, 1895). The ﬁrst descriptions of tyrosinase activity in mammalian tissue were by Gessard (1903), who reported that extracts of horse melanoma had the ability to convert tyrosine to melanin, and by Durham (1904), who described the presence of tyrosinase activity in mammalian skin. Hogeboom and Adams (1942) separated melanin biosynthesis into two enzymatic activities, the oxidation of tyrosine and the oxidation of dopa. Lerner et al. (1949) provided evidence that tyrosinase was responsible for both these

activities. Further details of the melanin biosynthetic pathway were reported by Mason et al. (1955). For many years, it was thought that tyrosinase was the sole enzyme involved in the biosynthetic pathway of melanin, which begins with the substrate tyrosine and produces either the characteristic brown/black eumelanin or red/yellow pheomelanin in the presence of sulfhydryl groups. Tyrosinase was known to be involved in the catalysis of the ﬁrst two steps, the oxidation of tyrosine to dopa and the subsequent formation of dopaquinone. All subsequent steps were thought to occur spontaneously without the need for other enzymatic catalysts. It was through the work of Logan and Weatherhead (1978), using Siberian hamsters, that other regulatory factors in the melanin biosynthetic pathway were shown to exist. They found that, during autumn molt, when the coat hair was unpigmented, tyrosinase activity was still present as well as normal serum tyrosine levels. They hypothesized that another factor must exist to prevent melanin from being formed. Previous to this, there had been suggestions of other regulatory factors in the melanin biosynthetic pathway, but this was the ﬁrst experimental evidence that such factors existed and affected coat color. Körner and Pawelek (1980) identiﬁed a factor, termed DOPAchrome conversion factor (DCF), in extracts of Cloudman S91 melanoma cells that could rapidly convert red/orange DOPAchrome to a colorless compound. Pawelek and coworkers also reported two other factors: 5,6-dihydroxyindole (DHI) conversion factor, which, when cells are exposed to melanocyte-stimulating hormone (MSH), converts DHI to indole-5,6-quinone, and DHI blocking factor, which stops melanin biosynthesis at 5,6dihydroxyindole (Pawelek et al., 1980). The DOPAchrome conversion factor was subsequently shown to be a new enzyme in the melanin biosynthetic pathway, and was termed DOPAchrome oxidoreductase and ﬁnally named DOPAchrome tautomerase (Aroca et al., 1990; Barber et al., 1984; Leonard et al., 1988). A third enzyme, 5,6-dihydroxyindole-2-carboxylic acid oxidase (DHICA oxidase), has also been included in the menagerie of enzymes involved in the melanin biosynthetic pathway. At present, no enzymes have been shown to be involved speciﬁcally in the pheomelanin pathway. As the complex enzymology of the melanin pathway was being worked out, attempts were being made to isolate gene sequences coding for these enzymes. Understandably, efforts to isolate pigment-speciﬁc genes began initially with the cDNA for tyrosinase. Instead of isolating the tyrosinase cDNA, a family of cDNAs coding for proteins related to tyrosinase was discovered. The ﬁrst “tyrosinase” cDNA clone isolated, 213

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termed pMT4, was later shown to code not for tyrosinase, but for a related protein termed tyrosinase-related protein (TRP1), the gene now known to code for DHICA oxidase (JiménezCervantes et al., 1994; Shibahara et al., 1986). This cDNA was found to be homologous to the brown (b) locus of the mouse (Jackson, 1988; Vijayasaradhi et al., 1990). Eventually, both the mouse and the human tyrosinase cDNA were cloned (Kwon et al., 1987; Yamamoto et al., 1987). The tyrosinase sequence was authenticated by comparison of the encoded amino acid sequence with results of direct amino acid sequencing of the isolated enzyme (Wittbjer et al., 1989) and by expression of the isolated tyrosinase cDNA in heterologous cells and observing tyrosinase enzymatic activity (Beermann et al., 1990; Bouchard et al., 1989; Müller et al., 1988). Two other related loci were identiﬁed and called the tyrosinaserelated protein-2 (TRP2), a product of the mouse slaty (slt) locus, shown to be the gene for DOPAchrome tautomerase (DT) (Jackson et al., 1992; Tsukamoto et al., 1992a), and a tyrosinase pseudogene (tyrosinase-related sequences; TYRL), which contains only exons 4 and 5 of the tyrosinase gene (Takeda et al., 1989a). In her analysis of tyrosinase in pigmented vertebrate skin, Durham (1904) also reported that tyrosinase activity was absent in albino animals, but she did not draw any conclusions from this observation. A hypothesized association between albinism in man and a lack of enzyme activity was ﬁrst made by Sir Archibald Garrod in his seminal paper entitled “Inborn Errors of Metabolism” (Garrod, 1908). In his report for the Croonian lectures, Garrod stated that “. . . an intracellular enzyme is probably wanting in the subjects of this anomaly . . . .” Garrod mentions tyrosinase as being the activity responsible for melanin pigment formation, but did not make a clear connection between a lack of tyrosinase activity and albinism. For many decades afterwards, it was not clear whether the lack of tyrosinase activity resulted from a mutation in the gene responsible for tyrosinase or the absence of melanocytes in the skin (Lerner and Fitzpatrick, 1950). Using gold impregnation to visualize melanocytes by light microscopy (Becker et al., 1952) and electron microscopy (Barnicot et al., 1955), melanocytes were identiﬁed in human albino skin and hair bulbs. Fitzpatrick et al. (1958), using the dopa reaction of Bloch (1917), later showed that albino hair follicles in both mouse and human skin contained melanocytes but lacked tyrosinase activity. The ﬁrst mutation within the tyrosinase gene associated with human oculocutaneous albinism (OCA) was reported in 1989 (OCA1; MIM #203199) (Tomita et al., 1989). This type of albinism, termed OCA1A, classically presents with a complete absence of pigment in the skin, eye, and hair, but some mutations produce a residual activity enzyme producing a phenotype with varying amounts of pigment (OCA1B) (Giebel et al., 1991a; King et al., 2000). Mutations of the tyrosinase-related protein-1 (TRP1) gene were shown to result in another type of albinism, OCA type 3 (OCA3, MIM #203290), also termed rufous or red OCA

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(MIM *278400; Boissy et al., 1996; Manga et al., 1997). Individuals with OCA3 are born with minimal pigmentation but develop an increase in pigment with time. The pigmentation changes in the skin, hair, and eyes, as well as the severity of the ocular involvement with OCA3, have not been as well documented as in other types of albinism, but the visual problems do not appear to be as severe in OCA3 as those observed in OCA1 (Kromberg et al., 1990). To date, there have been no reported human pigment phenotypes associated with mutations of TRP2.

Current Concepts Overview The formation of pigment in the melanocyte is a complex process requiring the action and interaction of many genes. In the mouse, there are over 100 loci that affect coat color (Bennett and Lamoreux, 2003; Silvers, 1979), and the majority of these loci are involved in melanocyte differentiation and migration. A subset of these loci is concerned with the formation of the premelanosome and biosynthesis and deposition of melanin in the premelanosome forming the mature melanosome. Three of these loci are members of a gene family that code for the enzymes, tyrosinase, TRP-1, and TRP-2, involved in the eumelanin biosynthetic pathway. Although these three proteins have different enzymatic activities, they share extensive homology in both nucleotide and amino acid sequences. The comparison of the promoter regions of these genes has provided an opportunity to locate common regulatory elements that may control the expression of pigmentspeciﬁc genes in a coordinated fashion. Detailed analysis of the 5¢ promoter region of these genes has indicated several possible cis-acting regulatory regions, some common to all three genes. Analysis of the proteins has extended our understanding of protein transport and processing, including the impact of missense mutations on this process.

Lower Vertebrates and Invertebrates Although the greater part of our understanding of the proteins and genes involved in pigmentation has come from studies involving the mouse and human pigment systems, other species have provided important insight into the evolution of the tyrosinase gene family. The earliest work with nonmammalian tyrosinase genes has been with mushroom tyrosinase. Other reports have included work with tyrosinase from the Streptomyces family and Neurospora crassa (Bernan et al., 1985; Huber et al., 1985; Katz et al., 1983). These systems have been instrumental in helping us understand the chemistry occurring in the active site of the enzyme. Lerch and colleagues have used site-directed mutagenesis to alter amino acids in the copper binding sites of Streptomyces tyrosinase to determine the amino acid residues critical to tyrosinase activity and their role in the catalytic site of the enzyme (Huber and Lerch, 1988; Jackman et al., 1991, 1992). These studies have shown

THE TYROSINASE GENE FAMILY

that histidine residues are the ligands involved in copper binding (Huber and Lerch, 1988; Jackman et al., 1991). Other amino acid residues such as the asparagine at codon 190 in Streptomyces glaucescens, adjacent to one of the histidine ligands in the copper B binding site, are thought to play a critical role in the hydrogen-bonding interaction that bridges the two copper atoms (Jackman et al., 1991). It was not until the mouse cDNA for tyrosinase was isolated that tyrosinase cDNAs from other vertebrates were cloned and their sequences determined. The isolation and sequence analyses of all three members of the tyrosinase gene family from the fugu ﬁsh (Takifugu rubripes) indicate the high degree of conservation of the pigmentary system in vertebrates (Camacho-Hubner et al., 2002). Partial sequences for all three members in Xenopus laevis have also been reported (Kumasaka et al., 2003). Other nonmammalian vertebrates in which tyrosinase sequences have been identiﬁed include the Japanese quail (Coturnix coturnix japonica), the snapping turtle (Trionyx sinensis japonicus), the Japanese pond frog (Rana nigromaculata), and goldﬁsh (Miura et al., 1995, Peng et al., 1995; Yamamoto et al., 1992). The high degree of nucleotide sequence homology between members of the tyrosinase gene family (see below) allowed Jackson and coworkers to create a series of degenerate oligonucleotide primers that ampliﬁed the cDNA of all three members of the tyrosinase gene family (Jackson et al., 1994). Using these ampliﬁcation primers, TRP1 cDNA from the Mexican axolotl (Ambystoma mexicanum) was isolated (Mason and Mason, 1995). These degenerate polymerase chain reaction (PCR) primer pairs should make it much easier to isolate the cDNA sequences of members of the tyrosinase gene family from other species. Numerous tyrosinase gene sequences are now available for all members of the tyrosinase gene family. Partial sequences for all three tyrosinase gene family members for many species can be found in the National Center for Biotechnology Information (Pruitt and Maglott, 2001). With the exception of the signal peptide sequence at the amino-terminal end, the amino acid sequences of these proteins are highly homologous.

The Tyrosinase Gene Family The tyrosinase gene family in humans consists of four loci, three of which produce protein products that are active in the melanin biosynthetic pathway. These include the enzymes tyrosinase (TYR; EC 1.14.18.1), DHICA oxidase (tyrosinaserelated protein-1; TRP-1), and DOPAchrome tautomerase (tyrosinase-related protein-2; TRP-2; EC 5.3.3.12). The fourth locus, the tyrosinase pseudogene (TYRL), is not expressed in melanocytes (Giebel et al., 1991b; Jackson et al., 1994; Takeda et al., 1991). Sequences from TYRL are found in humans and the gorilla, but are not found in other higher primates including the chimpanzee and Barbary ape (Oetting et al., 1993a), and are thought to be absent in lower vertebrates. No additional members are thought to exist in this gene family in humans. In the mouse genome, in addition to the Tyrp1 gene located on chromosome 4, there is also a

TYR TRP1 TRP2 Fig. 11.1. Comparison of the gene structures for tyrosinase, TRP1, and TRP2. The numbered boxes represent the exons of the gene. Vertical lines represent the location of intron/exon junctions. Vertical lines between genes represent identical or in-phase intron/exon junctions between those two genes.

tyrosinase-related protein-1-related sequence 1 (Tyrp1-rs1) on chromosome 8. Little is known about this locus, or whether it is an active gene or a pseudogene. Comparison of the genomic structure of the mouse and human homologs of tyrosinase, TRP1, and TRP2 genes shows that they have identical gene structure and splice junctions. Comparison of tyrosinase, TRP1, and TRP2 with each other, on the other hand, reveals numerous differences in the number of exons and the location of the splice junctions (Fig. 11.1). There are only two splice site junctions that are identical and one location that is similar. Comparisons between the splice sites for intron 2 of TYR and intron 5 of TRP1 and between the splice sites of intron 3 of TRP1 and TRP2 are identical (Budd and Jackson, 1995; Sturm et al., 1995). Intron 4 of TYR and intron 7 of TRP1 and TRP2 are not identical but have the same phasing. The intron/exon boundary and nucleotide sequences between TYR and TYRP are almost identical for exons 4 and 5 (Giebel et al., 1991b; Takeda et al., 1991). Both TRP1 and TRP2 are thought to have arisen from the TYR gene by means of gene duplication and subsequent divergence (Jackson, 1995a; Sturm et al., 1995). This event appears to have at least predated mammalian evolution, in that all mammals studied so far appear to have all three members of this gene family. Furthermore, all three genes have been found in Takifugu rubripes and Xenopus laevis, showing that all vertebrates may contain all three members (Camacho-Hubner et al., 2002; Kumasaka et al., 2003). As gene sequences of the tyrosinase gene family are isolated from more diverse species, direct comparisons of the gene structure should provide valuable insight into the evolution of this gene family. Sequence analysis of the cDNAs has shown that these genes have similar nucleotide sequences (30–50% similarity). Comparison of the amino acid sequences between TRP1 and TRP2 shows a closer relationship to each other (50% identify) than to TYR (40% identify) (Bouchard et al., 1994; Cassady and Sturm, 1994). Figure 11.2 shows the high degree of homology in the amino acid sequence between these three proteins. There are two consensus amino acid patterns that identify members of this protein family, as well as the hemocyanins, but no other proteins in the SWISSPROT database. These sequences are H-x(4,5)-F-[LIVMFTP]-x-[FW]-H-Rx(2)-[LVM]-x(3)-E associated with the copper A binding

215

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Fig. 11.2. Comparison of the amino acid sequence for tyrosinase, TRP1, and TRP2. The cysteine residues are shown in bold letters, and an asterisk has been placed over the histidine residues involved in copper binding. Vertical lines represent identical amino acids. Dashes represent gaps to aid in alignment of the sequences. The sequence for the EGF-like motif has been underlined.

216

THE TYROSINASE GENE FAMILY

Fig. 11.3. This represents similar structural domains between members of the tyrosinase family of proteins. The shaded box labeled “SP” is the signal peptide sequence. The dots represent cysteine residues. EGF is the location of the EGF-like domain. The hatched boxes Cu(A) and Cu(B) represent the copper-binding domains. The checkered box labeled TM represents the transmembrane domain.

domain and D-P-x-F-[LIVMFYW]-x(2)-H-x(3)-D associated with the copper B binding domain. In both cases, the “H” represents the histidine copper-binding ligands. Analyses of the amino acid sequences of these proteins reveal several regions of similarity that are potential functional domains. All three enzymes contain an amino-terminal signal sequence, 15 conserved cysteine residues, two putative copper-binding regions, a transmembrane region, and a cytoplasmic tail at the carboxy-terminal of the protein (Fig. 11.3) (Jackson et al., 1992; Sturm et al., 1995). These similarities, especially the conservation of the cysteine residues, present a strong indication that all three proteins have a common three-dimensional structure. The amino-terminal signal sequence has been shown to be important for correct processing of the tyrosinase polypeptide and its intracellular trafﬁcking (Oetting et al., 1995). Mutations that eliminate this portion of the protein for tyrosinase result in the enzyme being translated by cytoplasmic ribosomes instead of membrane-bound ribosomes. This results in an aberrantly folded protein that is quickly degraded (Oetting et al., 1995). The proteins also have two cysteine-rich domains (Figs 11.2 and 11.3). The ﬁrst cysteine-rich region also contains an EGF motif similar to that found in the EGF family, a motif that may be involved in protein–protein interactions (Jackson et al., 1992). This may be important in the formation of the melanin biosynthetic complex discussed later in the chapter. Although the copper(A) and copper(B) binding regions found in tyrosinase, including the copperbinding histidine ligands, can be identiﬁed in the other two proteins, copper has not been shown to be present in either TRP-1 or TRP-2. It has been hypothesized that TRP-1 and TRP-2 may have other metal ions as cofactors, but this has yet to be unambiguously established (Chakraborty et al., 1992; Halaban and Moellmann, 1990; Solano et al., 1994). The strongest evidence to date for binding of other metal cofactors has been binding of zinc by TRP-2 (Solano et al., 1994). Work by Furumura et al. (1998) showed that copper is bound most strongly by tyrosinase and less so by TRP-1. TRP-2 binds zinc and could also bind small amounts of iron, but TRP-1 did not bind zinc or iron. This difference in metal ligand binding is due, in part, to differences in the sequence in the metal-binding regions. It is expected that TRP-1 also has a metal cofactor, but the identity of this ligand has yet to be determined. The transmembrane region, found in all three proteins, is a highly hydrophobic region, as shown by hydrophobicity

mapping. Accordingly, all three proteins have been shown to be integral membrane proteins, anchored in the melanosomal membrane by the transmembrane domain (Orlow et al., 1994; Winder et al., 1994a). The carboxy-terminal cytoplasmic region has been shown to be important for the correct intracellular sorting and targeting of TRP-1 to the premelanosome and most likely plays an important role in the targeting of all three proteins. A series of experiments by Vijayasaradhi et al. (1995a) has shown the importance of the cytoplasmic tail region in intracellular sorting. A hexapeptide sequence, AsnGln-Pro-Leu-Leu-Thr, containing the dileucine motif ProLeu-Leu, is conserved among other melanosomal proteins including tyrosinase, Pmel-17, and the P gene product, but not TRP-2 or the melanocyte-stimulating hormone receptor (MSHR; Jimbow et al., 1995). This sequence has been identiﬁed as a sorting signal necessary for correct intracellular trafﬁcking of TRP-1 in melanocytic cells. This motif is also observed in some lysosomal membrane proteins, providing evidence for a similar sorting mechanism and pathways for both melanosomes and lysosomes, which will also include additional signals necessary to distinguish the movement of these proteins between the two organelles. The importance of the transmembrane region for correct protein function has also been shown in vivo by two frameshift mutations in the human tyrosinase gene. A frameshift mutation at codon 490 eliminates one half of the transmembrane region and the entire cytoplasmic tail, and a second frameshift mutation at codon 501, located at the carboxy end of the transmembrane region, effectively eliminates the cytoplasmic tail (Chintamaneni et al., 1991a; Giebel et al., 1991a; King and Oetting, 1992; Tripathi et al., 1992a). In both cases, these mutations result in the complete absence of enzymatic activity and are associated with tyrosinasenegative oculocutaneous albinism (OCA1). A mutation in the mouse tyrosinase gene responsible for the platinum phenotype also causes a truncated protein that lacks the cytoplasmic tail (Beermann et al., 1995; Orlow, 1995; Orlow et al., 1993a). The lack of an intact cytoplasmic tail due to the platinum mutation results in the tyrosinase protein being routed to the plasma membrane rather than to the melanosome, causing a severe reduction in tyrosinase activity. The transmembrane region and the cytoplasmic tail appear only to be involved in the correct targeting of the enzyme and do not play a role in the catalytic function of the enzyme. Tryptic cleavage of the mature enzyme, which removes the transmembrane region and the cytoplasmic tail, leaves the truncated enzyme catalytically active.

Tyrosinase Gene/Tyrosinase Tyrosinase is a copper-containing protein and is the rate-limiting enzyme in the melanin biosynthetic pathway. Tyrosinase catalyzes two major steps in melanin formation, the hydroxylation of tyrosine to DOPA, which is the rate-limiting step, and the oxidation of DOPA to DOPAquinone. The catalytic site of the enzyme contains two copper atoms ligated to six 217

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histidine residues. The hydroxylation of tyrosine apparently begins with the binding of dioxygen to the Cu1+ Cu1+ center of tyrosinase to yield a peroxide-bound Cu2+ Cu2+ center termed oxytyrosinase (Lerch, 1988; Mason et al., 1955; Wilcox et al., 1995). It has been proposed that a tyrosine hydroxylate complex forms with one copper of the cluster placing the 3 carbon of the substrate in position to accept the terminal oxygen of the peroxide, which is bound to the second copper to form DOPA. This o-diphenol is then oxidized to the quinone (DOPAquinone) by the enzyme and, upon release, regenerates the Cu1+ Cu1+ center. It is thought that tyrosinase folds in the presence of the protein chaperone calnexin, allowing trimming of sugar chains by a-glucosidase and copper binding (Branza-Nichita et al., 1999; Jimbow et al., 1994). Proper folding of tyrosinase also appears to require other proteins, such as TRP1 and the substrate tyrosine (Francis et al., 2003). Tyrosinase is then transferred from the Golgi network to the premelanosome via interaction between the dileucine motif and a tyrosine-based signal and the AP3 adaptor-like complex (Blagoveshchenskaya et al., 1999; Höning et al., 1998; Le Borgne et al., 1998; Simmen et al., 1999). The tyrosinase gene (TYR) maps to human chromosome 11q14 (Barton et al., 1988). The tyrosinase cDNA codes for a 529-amino-acid polypeptide with a predicted molecular weight of 55 kDa and a mature processed glycoprotein protein of 75 kDa. TYR contains ﬁve exons spanning 118 kb (Giebel et al., 1991b; Ponnazhagan et al., 1994). The sizes of the introns are 12.4 kb for intron 1, 36.4 kb for intron 2, 56.8 kb for intron 3, and 10.2 kb for intron 4 (Pruitt and Maglott, 2001). Over 50% of the coding region is found in exon 1 (819 bp) with exons 2 to 5 ranging from 148 to 225 bp. The tyrosinase gene has been shown to produce many alternatively spliced transcripts, but their function, if any, is unknown (Fryer et al., 2001). The tyrosinase gene is the albino (c) locus of the mouse. Mutations that completely eliminate tyrosinase enzymatic activity produce a mouse with a complete absence of melanin pigment in the coat and eye. Several mouse tyrosinase alleles have been isolated that result in a spectrum of pigment loss from a complete absence of pigment (the null mutations) to moderate amounts of pigment including a temperaturesensitive mutation (ch) (Beermann et al., 1990; Jackson, 1995b; Jackson and Bennett, 1990; Kwon et al., 1989; Orlow et al., 1993a; Porter et al., 1991; Schmidt and Beermann, 1994; Shibahara et al., 1990; Silvers, 1979; Yokoyama et al., 1990). Of all the gene products that participate in pigmentation at the subcellular level of the melanocyte, mutations at the c locus have the greatest effect on the reduction in pigmentation. Numerous mutations of the human tyrosinase gene associated with tyrosinase-related OCA have been reported (Oetting and King, 1999). The 5¢ upstream promoter region of the human TYR gene contains four transcriptional start sites with the major site –79 bp from the ATG start codon and minor sites at –78, –42, and –34 bp (Ponnazhagan et al., 1994). Other transcriptional 218

start sites have been reported and may represent differences in the cell lines used (Giebel et al., 1991b; Kikuchi et al., 1989; Takeda et al., 1989b). There are also several potential proximal cis-regulatory elements including a putative TATA box (–106 bp) and CAATT box (–199 bp). Another potential regulatory element is a 230-bp repeat sequence of (GA:TC)n located 713 bp upstream of the translational start site, which can assume a hinged-DNA structure and may play a role in regulation of tyrosinase expression (Kikuchi et al., 1989). This repeat sequence in humans has been found to be polymorphic in several different populations (Morris et al., 1991; Oetting et al., 1993b). There are also numerous AP-1, AP-2, and Oct1 binding cis-acting elements (Ponnazhagan et al., 1994). In an effort to understand the transcriptional control of tyrosinase, there have been several studies to determine the number and location of cis-acting regulatory elements and the trans-acting factors that bind to sequences necessary for both temporal and tissue-speciﬁc expression. In expression studies with the mouse tyrosinase gene, as little as 270 bp of the 5¢ promoter has produced cell-speciﬁc and temporal-speciﬁc expression in transgenic mice, although wild-type levels of pigmentation were not achieved (Beermann et al., 1992; Kluppel et al., 1991). In an effort to mimic wild-type expression, transgenic mice were produced using a 250-kb yeast artiﬁcial chromosome (YAC) containing 155 kb of the 5¢ ﬂanking sequence (Schedl et al., 1993). In these experiments, wild-type levels and distribution of pigmentation were observed in a copy numberdependent and position-independent manner, suggesting that in vivo tyrosinase gene expression is regulated by cisregulatory elements located as far upstream as 155 kb. Individual elements involved in the regulation of the tyrosinase gene have now been identiﬁed. An element –183 bp upstream of the start codon, TATCAATTAG, was found to bind the transcription factor HMG-1 (high mobility group protein) and its isoform HMG-Y (Sato et al., 1994). HMG1(Y) is thought to be involved in nucleosome phasing, a mechanism for maintaining the undifferentiated state of chromatin. Possibly, this protein alters the structure of the promoter or facilitates the binding and activity of other trans-acting factors at the tyrosinase promoter. At present, the binding of HMG1(Y) has only been shown by in vitro studies and further work is needed. For the human TYR locus, a region between –2020 and –1739 bp from the transcriptional start site was found to contain an important enhancer (Ponnazhagan et al., 1994; Shibata et al., 1992a). The enhancer sequences were localized to a 39-bp core element that was conﬁrmed to direct the melanoma cell-speciﬁc expression of a reporter gene under the control of this element (Shibata et al., 1992a). This region was later termed the tyrosinase distal element (TDE) (Yasumoto et al., 1994). The TDE contains one of four copies of a 6-bp motif, CATGTG, found in the 5¢ promoter of the tyrosinase gene, which is also present in the mouse tyrosinase promoter. This same 6-bp motif is found at –104 bp to –99 bp and has been labeled as the “M”-box or the tyrosinase proximal element (TPE) (Lowings et al., 1992; Yasumoto et al., 1994).

THE TYROSINASE GENE FAMILY

The M-box element has also been found in the 5¢ promoter region in the mouse and human TRP1 and TRP2 genes and is considered to be an important cis-regulatory element for melanocyte-speciﬁc expression (Hemesath et al., 1994; Jackson et al., 1991; Lowings et al., 1992; Shibahara et al., 1991; Sturm et al., 1995). The third location of this element at –12 bp to –7 bp has been termed the “E”-box (Bentley et al., 1994). Expression studies have shown that the TDE is the strongest of these three elements, with the M-box element being much weaker (Yasumoto et al., 1994). Other elements, both positive and negative, have been reported for the tyrosinase gene (Bentley et al., 1994). The CANNTG motif binds a family of transcription factors, characterized by the presence of a conserved helix–loop–helix structure required for DNA binding and activation (Hodgkinson et al., 1993). It is now established that the protein product of the human homolog of the mouse microphthalmia (mi) gene, a melanocyte-speciﬁc factor called the microphthalmiaassociated transcription factor (MITF), binds to both the TDE and the M-box regions (Hemesath et al., 1994; Yasumoto et al., 1994). The murine mi gene product has been shown to increase tyrosinase and TRP1 transcription as well as reporter genes with the M-box in the promoter. The human MITF gene was found to direct transcription of both the mouse tyrosinase and TRP1 genes preferentially in melanocytic cells (Yasumoto et al., 1995). Yasumoto et al. (1995) showed that an 82-bp cis-acting region, which contains the CATGTG motif (position –12 to –7) along with the M-box (positions –107 to –97), was involved in transactivation of the tyrosinase gene in vitro, and concluded that the trans-acting factor MITF is a common factor regulating transcription of the pigment cell-speciﬁc genes. A second factor, termed a “ubiquitous transcription factor,” was also found to bind the M-box, but its role in activating melanocyte-speciﬁc transcription is unknown (Lowings et al., 1992). Other transcription factors, N-Oct-3 and NOct-5, are expressed at high levels in melanoma cells and may be involved in mechanisms controlling the transcription of pigment-speciﬁc genes (Sturm et al., 1994a). Another cis-acting regulatory region has been found –15 kb from the initiation codon (Porter et al., 1991; Porter and Meyer, 1994). In melanoma cells, but not in other nonpigmenting cell types, this region was shown to contain two DNase I hypersensitive sites (HS-site) within a 200-bp region (Ganss et al., 1994). These two DNase I hypersensitive sites were found to be rearranged in the chinchilla-mottled (cm) mouse, resulting in hypopigmentation, showing the importance of these sites for the proper transcription of tyrosinase (Porter et al., 1991). The ﬁrst site, HS-site (hs1), contained the palindrome TGACTTTGTCA, which resembles the binding motif to both a CREB (cAMP-responsive element binding) transcription factor and the AP-1 (activator protein-1) binding site (Ganss et al., 1994). The inclusion of this region into transgenic mice enhanced tyrosinase expression (Porter and Meyer, 1994). Interestingly, tyrosinase expression was enhanced in neural crest-derived melanocytes but not in neuroectoderm-derived melanocytes, indicating differential

regulation of tyrosinase expression in these two cell lineages (Porter and Meyer, 1994). A similar region has been identiﬁed in humans –9 kb upstream of the transcriptional start site (Fryer et al., 2003; Regales et al., 2003). The homologous human sequence was initially found using DNA sequence comparisons with the upstream region of the mouse tyrosinase gene. DNase analysis showed that these sequences exhibited DNase hypersensitivity. This region was cloned and placed upstream of the proximal tyrosinase promoter and a luciferase reporter gene. The core sequences of this region were shown to speciﬁcally increase reporter expression in melanocytes in conjunction with the tyrosinase proximal promoter (Fryer et al., 2003; Regales et al., 2003). Preliminary evidence has shown that this region may also contain a negative regulatory region. The trans-acting factors that interact with these sequences still need to be determined. These reports show that the regulation of tyrosinase gene expression is complex and requires the coordinated participation of several negative and positive regulatory elements and factors. As shown above, these regulatory elements are both proximal (e.g. the E-box; –6 bp) and distal (e.g. hs-site; 12–15 kb). Melanocyte-speciﬁc and temporal regulation of tyrosinase gene expression is thus achieved by a combination of trans-acting factors that bind to these regulatory elements.

TRP1 Gene/DHICA Oxidase The tyrosinase-related protein-1 (TRP1), the second member of the tyrosinase gene family, remains an enigmatic protein. Accordingly, there have been several different enzymatic functions attributed to DHICA oxidase, including tyrosine hydroxylation (an enzymatic activity of tyrosinase) (Jiménez et al., 1991; Tsukamoto et al., 1992b), DOPAchrome tautomerase (Winder et al., 1994b), and catalase activity (Halaban and Moellmann, 1990). The last activity was used to ascribe the locus name of CAS2 to this gene. Current information shows that the puriﬁed protein has DHICA oxidase activity and places it distal to tyrosinase and DOPAchrome tautomerase in the melanin biosynthetic pathway (Jiménez-Cervantes et al., 1994). Interestingly, despite the high degree of homology with the mouse homolog, human TRP-1 protein does not appear to have DHICA oxidase activity (Boissy et al., 1998). The gene (tyrosinase-related protein-1; TRP1) maps to human chromosome 9p23 (Abbott et al., 1991; Chintamaneni et al., 1991b; Murty et al., 1992). The TRP1 gene consists of eight exons within a 25-kb region, with exons 2 through 8 containing the coding sequences (Jackson et al., 1991; Shibahara et al., 1991; Sturm et al., 1995). The TRP1 gene encodes a polypeptide of 537 amino acids with predicted molecular weight of 60 kDa (Cohen et al., 1990; Urquhart, 1993). The mature glycosylated protein has, like tyrosinase, an apparent molecular weight of 75 kDa. In the mouse genome, there is an additional DNA segment that cross-hybridizes to TRP1 probes (Jackson, 1988; Shibahara et al., 1991). This segment contains sequences homologous only to exon 8 of TRP1 (Shibahara et al., 1991). The sequences between the crosshybridizing segment and exon 8 or TRP-1 are 95% identical, 219

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but the segment contains a single base deletion, resulting in a premature stop codon, and is most likely a nonfunctional pseudogene. This is most likely the Tyrp1-rs1 locus. At present, there are no reports of a TRP1 pseudogene in any other species. The brown (b) locus of the mouse encodes the murine TRP1 gene. Mutations at this locus result in the production of brown eumelanin instead of black eumelanin. There are several known alleles of the b locus (Jackson, 1995b; Jackson et al., 1990; Johnson and Jackson, 1992; Silvers, 1979; Zdarsky et al., 1990) as well as several artiﬁcially generated deletions that include the b locus (Bell et al., 1995). The recessive mutations result in loss of function in the homozygous state, varying the degree of brown melanin pigment produced in an otherwise normal melanocyte. Two mutations, Light (Blt) and White-based brown (Bw) are dominant mutations and result in melanocyte death (Jackson, 1995b; Johnson and Jackson, 1992). The Light mutation produces a protein with aberrant function that is thought to disrupt the melanosome, allowing toxic intermediates to enter the cytoplasm of the cell, causing cell death. The White-based brown mutation has a chromosomal inversion that places the TRP1 promoter in another chromosomal region. This results in the inactivation of the TRP1 gene and most probably the activation of another protein that is responsible for the observed cell death. Mutations of the human TRP1 gene have been shown to be responsible for brown OCA (OCA3) (Boissy et al., 1996, Manga et al., 1997). Sequences in the ﬁrst exon of the TRP1 gene appear to be important in determining the efﬁciency of its expression, but are not sufﬁcient to confer pigment cell-speciﬁc expression (Shibata et al., 1992b). The ﬁrst intron was shown to be necessary for TRP1 transcription in transient transfections. Removal of the ﬁrst intron, even in the presence of 3 kb of the upstream TRP1 promoter region, reduced expression to background levels (Sturm et al., 1994a). Other reports have shown that the ﬁrst intron is not necessary for TRP1 expression in cultured cells (Jackson et al., 1991; Lowings et al., 1992). The transcriptional initiation site is 20 bp downstream of the TATAAA box (Sturm et al., 1994a). The minimum promoter in the mouse gene is between –44 bp and +107 bp from the start codon. Within this minimal promoter, both positive and negative regulatory elements have been identiﬁed (Lowings et al., 1992; Yamamoto et al., 1992). The M-box regulatory element is one of these elements, which is also found in the tyrosinase gene. The 5¢ upstream region of TRP1 was found to bind to regulatory factors of 50 kDa and 43 kDa (Yamamoto et al., 1992). One of these factors binds to the Mbox element and acts as a positive regulatory factor.

TRP2 Gene/DOPAchrome Tautomerase The tyrosinase-related protein-2 (TRP2) is the third member of the tyrosinase gene family and codes for an enzyme with DOPAchrome tautomerase (DT) activity. TRP-2 maps to human chromosome 13q32 (Sturm et al., 1994b; Bouchard et al., 1994). This gene was initially isolated from an expressed 220

mouse melanoma cDNA library using rabbit antimouse tyrosinase antibody and rabbit antihamster tyrosinase antibody and designated as clone 5A (Jackson, 1988; Jackson et al., 1992). This clone was found to map to the slaty (slt) locus of the mouse and shown to code for DT using an expression vector containing the DT cDNA and expression in COS cells (Kroumpouzos et al., 1994; Tsukamoto et al., 1992a; Yokoyama et al., 1994). The human cDNA for DT was also isolated and sequenced (Bouchard et al., 1994; Cassady and Sturm, 1994; Yokoyama et al., 1994). The coding sequences of the mouse and human TRP2 genes are divided into eight exons spanning a 55-kb region in humans with the translational start site in exon 1 (Budd and Jackson, 1995; Sturm et al., 1995). The increased size of the gene, compared with TRP1, is due to an increased number of repetitive elements. In the mouse, several alleles of the slaty locus have been identiﬁed (Budd and Jackson, 1995; Green, 1972; Jackson, 1995b; Jackson et al., 1992). Mutations at this locus result in a dilution of the coat color and slightly yellowish ears (Green, 1972). One mutation, slaty light, acts as a semi-dominant mutation. None of these alleles is thought to be a null allele. To determine the phenotype of a known DT null mouse, a TRP-2 knockout mouse was produced (Guyonneau et al. 2004). The knockout mice were found to be viable and exhibited a diluted coat color phenotype with no detectable abnormalities beyond hypopigmentation in DT-expressing sites such as skin, retinal epithelium, or brain. The phenotype was identical to the slaty mutation. Moreover, unlike slaty mutant melanocytes, DT knockout primary melanocytes were much easier to grow in culture (Guyonneau et al. 2004). At present, no mutations in the human TRP2 gene have been reported. The human DT amino acid sequence has 83% identity and 90% similarity to the mouse sequence (Cassady and Sturm, 1994). Northern blot analysis has shown DT to be expressed as a 4.5-kb and 2.3-kb transcript, whereas the mouse only produces a 2.3-kb transcript (Bouchard et al., 1994; Yokoyama et al., 1994). HeLa cells, which have no endogenous DT activity, exhibited DT activity when transfected with the human DT cDNA, similar to COS cells transfected with the mouse DT cDNA, showing that the TRP2 gene codes for dopachrome tautomerase. Analysis of several different human melanoma cell lines revealed different levels of expression, ranging from no expression through low to moderate to high levels of expression (Bouchard et al., 1994; Cassady and Sturm, 1994). Normal human melanocytes were found to express DT at moderate levels but, in some cell lines, levels of enzymatic activity did not correspond to transcript levels, suggesting that there may be a post-transcriptional regulatory mechanism. There was no direct correlation between the expression of TRP2 and the presence of visible melanin and/or activity of tyrosinase (Bouchard et al., 1994). Like tyrosinase, TRP2 contains the M-box element that binds the MITF transcription factor. It has been shown that SOX10 also binds promoter elements in TRP2 (Jiao et al., 2004). Sox10 binding was demonstrated using gel shifts of

THE TYROSINASE GENE FAMILY

oligonucleotide probes derived from promoter sequences within the region required for Sox10-dependent induction. It was shown that Sox10 acts directly upon the Dct promoter to activate gene expression (Jiao, et al., 2004). It was further hypothesized that Sox10 and Mitf synergistically activate TRP2 expression.

Transcriptional Regulation It has been known for many years that there is a direct response of the pigment system in melanocytes to speciﬁc factors such as melanocyte-stimulating hormone (MSH) and UV irradiation. For this general increase in pigment production, it has been assumed that there is an increase in the expression of all pigment genes involved in melanin biosynthesis and premelanosomal formation. Several common regulatory elements for tyrosinase, TRP1, and TRP2 have been identiﬁed, and these may be involved in a coordinated regulation of these genes (Shibahara et al., 1991; Sturm et al., 1995; Yamamoto et al., 1992). One regulatory element found to be shared by all three members of the tyrosinase gene family is the M-box (Jackson, 1994). This element is conserved between mouse and human upstream regions in all three members of the tyrosinase gene family, as well as in regulatory elements much more distal (Shibata et al., 1992a; Sturm et al., 1995). One binding factor, termed the M-box binding factor, binds to the M-box element and acts as a positive regulatory factor (Lowings et al., 1992). This is most likely the microphthalmia-associated transcription factor (MITF; Tassabehji et al., 1994). Mutations of MITF result in microphthalmia in the mouse and Waardenburg syndrome type II in humans, which includes alteration in pigmentation due to defects in melanocyte migration. It should be noted, however, that transcriptional regulation of tyrosinase, TRP1, and TRP2 is not always as coordinated as one might expect. Molecular analysis has revealed a high degree of complexity in the transcriptional regulation of these genes. Using mouse B16-F1 cells, it was found that transcription of tyrosinase and TRP1 is coordinately regulated by melanotropic reagents via cAMP-dependent protein kinase and protein kinase C (Kuzumaki et al., 1993). Melanotropic reagents that increased tyrosinase transcription (i.e. cholera toxin, a-MSH, and dibutyryl cAMP) also increased TRP1 transcription. Reagents that decreased eumelanin synthesis, such as 12-O-tetradecaylphorbol 13-acetate (TPA), decreased mRNA levels of both tyrosinase and TRP1. This increase in tyrosinase and TRP1 mRNA resulted in a parallel increase in eumelanin production. Another study showed that this connection between TRP1 protein levels and pigment production occurs only in cells that are producing eumelanin and not in pheomelanin-synthesizing cells (del Marmol et al., 1993). In cells that are producing only pheomelanin, TRP1 mRNA is absent, whereas tyrosinase mRNA is present in all cells producing pigment regardless of the type of melanin being produced. This is not unexpected because TRP1 is thought to be involved only in the eumelanin biosynthetic pathway and not in the pheomelanin pathway. This is also shown by the

fact that murine TRP1 (brown locus) mutations do not affect the coat color of agouti lethal yellow (Ay) or extension (e) mice, which produce only pheomelanin (del Marmol et al., 1993), revealing that there must be different cis- and transacting transcriptional regulators for tyrosinase and TRP1. In other reports, the addition of 2% dimethyl sulfoxide (DMSO) or 5 mM hexamethylenebisacetamide (HMBA) on melanoma cell lines inhibited the expression of TRP1 but had no effect on tyrosinase expression (Sturm et al., 1994a). Vijayasaradhi et al. (1995a, b) found that HMBA reduced the expression of TRP1 to 88% of control but, for several cell lines, tyrosinase expression increased. Sturm et al. (1994a) also found that the addition of differentiating chemicals to melanoma cell lines exhibited opposing effects on tyrosinase and TRP1 expression (Martínez-Liarte et al., 1992). Selective regulation of TRP1 appears to be mediated by protein factors that modulate MITF-mediated transcription (Fang et al., 2002). TRP2 (DOPAchrome tautomerase) also exhibits expression patterns that differ from those of tyrosinase. Addition of a-MSH or other melanogenic activators has opposite effects on tyrosinase and DOPAchrome tautomerase activities (Martínez-Liarte et al., 1992; Sturm et al., 1994a). The addition of a-MSH on B16/F10 melanoma cells caused an increase in tyrosinase activity but a decrease in DOPAchrome tautomerase activity. In another study, Bouchard et al. (1994) found no direct correlation between the expression of TRP2 and pigment production in 10 different human melanoma cell lines. In some cell lines, TRP2 transcripts correlate with DT activity, but there were variations in others. Furthermore, there was no direct relationship between the presence of eumelanin and DOPAchrome tautomerase activity. Moreover, no consistent direct relationship between levels of tyrosinase activity and the amount of visible pigment have been found, suggesting that post-transcriptional and/or -translational regulatory mechanisms play an important role in the control of pigment production (Bouchard et al., 1989, 1994). One possible element interacting at the post-translational level is an inhibitor of these enzymes (Kameyama et al., 1993). One inhibitor isolated from the human amelanotic melanoma cell line SK-MEL-24 was found to suppress the oxidation of DHI in the presence or absence of tyrosinase, but had no effect on DHICA oxidation (Kameyama et al., 1995). Many of these reports analyzing the expression of tyrosinase, TRP1, and TRP2 show conﬂicting results. This is in part due to laboratories using different cell lines that produce different amounts and types of melanin pigment (eumelanin, pheomelanin, or both) (Bouchard et al., 1994; del Marmol et al., 1993; Taniguchi et al., 1992). It is important that the amount and type of melanin being produced be taken into account when the expression and activity levels of these genes are being determined. Although melanoma cells are convenient model systems to study pigmentation, it must be borne in mind that malignant transformation itself may disrupt the ﬁne balance of the activities of pigmentation machinery. Primary cultures of melanocytes may represent a system closer 221

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to the physiological system, although primary cell lines may still not be an accurate representation of what is happening in the melanocyte in situ. There are also differences in gene expression during embryonic development. Steel et al. (1992) found that DT is expressed very early in developing mouse embryos, 10 days post coitum, whereas TRP1 is expressed at 11.5 days post coitum and tyrosinase transcripts are observed 13.5 days post conception.

Protein Interactions Tyrosinase, TRP-1, and TRP-2, along with the lysosome-associated membrane protein-1 (LAMP-1), have been shown to interact as a high-molecular-weight multimeric enzymatic complex that ranges in size from 200 to >700 kDa (Hearing et al., 1982; Orlow et al., 1993b, 1994; Zhou et al., 1993). This complex is responsible for the conversion of tyrosine to eumelanin but does not appear to be involved in pheomelanin biosynthesis. Copuriﬁcation of these proteins with antityrosinase antibodies suggests a stable interaction between these proteins in which the EGF motif may be important in the formation and stabilization of this complex (Jackson et al., 1992). Recent studies have shown that tyrosinase activity is stabilized in the presence of TRP-1 and TRP-2, providing further evidence of a functional melanogenic complex (Pawelek, 1991; Winder et al., 1994a). Cotransfection of ﬁbroblast with both tyrosinase and TRP-1 showed an increase in both the stability and the activity of tyrosinase with time. This complex can interact with other factors as well, including melanogenic inhibitors (Kameyama et al., 1989) and products of other pigment loci that determine the quantity and quality of melanin pigment synthesized, including the silver (si), pink-eye (p), and MART-1 proteins (Kameyama et al., 1993). Analysis of mice with mutations at different pigment loci can provide information on the effects of protein interaction in vivo. By assaying melanin production in mouse skin, it was shown that mutations at the pink-eye loci will reduce tyrosinase activity (Coleman, 1962) as well as reduce the levels of tyrosinase transcript in mouse melanocytes compared with normal controls (Tamate et al., 1989). More recently, it has been shown that the pink-eye unstable (Pun) mutation, a mutation that contains an integral duplication inside the P gene resulting in altered P-gene transcription, actually disrupts the tyrosinase enzyme complex (Chiu et al., 1993). In the ocular melanocytes of mice that contained the Pun mutation, the highmolecular-weight enzyme complex could not be detected, and protein levels of tyrosinase, TRP1, and TRP2 were drastically reduced. Another example of the disruptive effect of a mutation in one of the members of this complex is found in the platinum (cp) mouse (Orlow et al., 1993a). This mutation of the mouse tyrosinase gene results in a truncated protein and a coat almost completely devoid of pigment. Enzymatic activity of platinum tyrosinase is higher than that of other tyrosinase mutations that result in a darker coat color, but the tyrosinase enzymatic complex cannot be detected, suggesting 222

that the absence of the complex causes a more severe form of albinism than would be expected by the tyrosinase gene mutation alone. More importantly, alteration of one of these protein products by means of mutations can have a drastic effect on other proteins involved in this complex. It is possible that mutations that result in hypopigmentation (or hyperpigmentation) may not act by speciﬁcally altering the function of the protein (i.e. reduced catalytic activity), but by disruption of the protein complex, producing a reduction in pigment biosynthesis. In addition to the potential interactions among family members that affect pigmentation, individual tyrosinase family proteins, speciﬁcally their cytoplasmic domains, also interact with components of the intracellular protein sorting machinery (reviewed by Setaluri, 2000). Although all tyrosinase family proteins follow a similar biosynthetic route to melanosomes and are expected to share interactions with the same components of trafﬁcking machinery, it is of interest to note that differences exist in their binding speciﬁcities. For example, whereas interaction of tyrosinase with the adaptor protein AP-3 is required for its trafﬁcking, interaction with AP-3 is not necessary for TRP-1 trafﬁcking (Huizing et al., 2001). Similarly, TRP-1, but not tyrosinase, interacts speciﬁcally with a multifunctional PDZ-domain protein, GIPC, during its transport through the Golgi (Liu et al., 2001).

Human Pigmentation Disorders Oculocutaneous albinism (OCA) is the most common human genetic disease associated with mutations of pigment genes that act at the subcellular level of the melanocyte. OCA is a clinically heterogeneous genetic disease. Mutations in numerous genes including tyrosinase (OCA1), P (OCA2), TRP-1 (OCA3), membrane-associated transporter protein (MATP; OCA4), Chediak–Higashi syndrome (CHS), and Hermansky–Pudlak syndrome (HPS1–7) are thought to be responsible for different forms of OCA in humans. Although, in certain strains of mice, mutations in TRP-1 have also been found to be responsible for a form of pigmentary glaucoma involving iris stromal dystrophy, no TRP-1 mutations were found to be associated with this disease in humans (Anderson et al., 2002; Lynch et al., 2002). Based on the observation that administration of L-DOPA to cytochrome P450 and TYR null mice alleviates developmental ocular abnormalities found in this model for human primary congenital glaucoma (PCG), it is proposed that the tyrosinase/L-DOPA pathway modiﬁes human PCG (Libby et al., 2003). Because of the effect of ocular albinism gene 1 (OA1) on cutaneous melanosome development, this locus may also be considered to be associated with a type of OCA. Besides their role in melanin pigmentation and pigmentary disorders, tyrosinase family proteins are also involved in the biology of melanoma. For example, levels of expression of TRP-2 in melanoma cells appear to correlate positively with their resistance to certain cancer chemotherapeutic drugs (Chu et al., 2000). The exact mechanism by which TRP-2 confers drug resistance is not known. Paradoxically, elevated

THE TYROSINASE GENE FAMILY

expression of TRP-2, and also tyrosinase, in advanced-stage metastatic melanoma lesions appears to correlate with improved overall survival of patients (Takeuchi et al., 2003). One possible reason for this may be the fact that, in patients with metastatic melanoma, all three proteins are recognized by the immune system (Brichard et al., 1993; Vijayasaradhi et al., 1990; Wang et al., 1995, 1996). Accordingly, a number of strategies are being evaluated, in clinical trials, to elicit immune response to these antigens for immunotherapy of melanoma (Rosenberg, 2001). Much of our understanding about human albinism has come from the work done on mutations of the tyrosinase gene and their relationship to tyrosinase-related OCA (OCA1). This relationship has been shown by both formal linkage of the tyrosinase gene to OCA1 and the ﬁnding of mutations in the tyrosinase gene that eliminate tyrosinase enzymatic activity (Giebel et al., 1990). Using a combination of PCR ampliﬁcation of individual exons and single-strand conformational polymorphism (PCR-SSCP) analysis, a number of mutations have been found (Oetting et al., 1994; Oetting and King, 1999). Mutations that produce a tyrosinase protein completely devoid of enzymatic activity are associated with the severest form of albinism, OCA1A. The missense mutations, on the other hand, result in a spectrum of phenotypes ranging from a complete absence of pigment production to near normal amounts of pigment. These latter mutations have residual amounts of enzymatic activity that allow a small amount of pigment to be produced. The missense mutations of tyrosinase have provided a tool to help us understand the biology of the enzyme and pigment production. At present, the three-dimensional structure of tyrosinase family proteins is unknown, and the effects of different mutations cannot be analyzed at the level of protein structure. However, analysis of mutations and their effect on enzymatic activity have provided some insight into the structure of tyrosinase. Mapping of the missense mutations associated with OCA1 reveals several clusters within the tyrosinase polypeptide. This clustering is thought to deﬁne functional domains of the enzyme (King et al., 1991; Oetting and King, 1992a, b; Tripathi et al., 1992b). Two of these domains are within the putative copper(A) and copper(B) binding sites respectively. A third cluster is at the 5¢ end (amino-terminal end of the polypeptide) of the coding region. This latter domain may deﬁne a putative substrate and/or cofactor binding site, or may be involved in enzyme stability. Missense mutations within the signal peptide of tyrosinase have allowed us to analyze the importance of this region in the enzymatic activity (Breimer et al., 1994; Fukai et al., 1995). Tyrosinase molecules that lack the signal peptide were found to be translated on free ribosomes instead of membranebound ribosomes. This resulted in a peptide that is incorrectly folded and quickly eliminated by the cell (Oetting et al., 1995). Frameshift mutations in the carboxy-terminal end of the protein also result in an enzyme without enzymatic activity. This region contains the transmembrane region and the cytoplasmic tail, which is important for proper sorting and

melanosomal targeting of these proteins (Vijayasaradhi et al., 1995a). The copper binding site of hemocyanin has many similarities to that of tyrosinase (Beltramini et al., 1990). Determining how different mutations affect tyrosinase activity can be aided using the structure of the copper-containing protein hemocyanin in which the three-dimensional structure has been determined (Gaykema et al., 1984, 1986). Based on amino acid sequence alignment between the copper-containing protein hemocyanin and tyrosinase, and extensive spectroscopic studies of tyrosinase, it is evident that the ligation of the copper atoms involves histidine side-chains from the protein. Site-directed mutagenesis of Streptomyces glaucescens tyrosinase has shown that the predicted histidine ligands are indeed involved in copper binding (Lerch, 1988). The presence and placement of these histidine ligands is likely to be critical for the binding of the copper atoms, which is necessary for both the formation of the reactive peroxide species and the complex with the substrate. Thus, changes in the amino acids that form the copper ligands or that change their position in the protein structure are likely to alter the rate and/or speciﬁcity of tyrosine hydroxylation. By using hemocyanin as a model for the copper binding sites of tyrosinase, we can begin to determine possible mechanisms of how some of these tyrosinase mutations cause inactivation of or a reduction in enzymatic activity (Oetting and King, 1994). The predicted secondary structure at both the copper binding sites of tyrosinase indicates that the histidine ligands responsible for copper binding are within two a-helices connected by a loop region (Oetting and King, 1992b). This same type of structure exists in the copper binding sites of hemocyanin (Gaykema et al., 1984, 1986). The amino acid sequences of the copper(B) binding region for hemocyanin and tyrosinase are very similar, but the copper(A) binding regions for both proteins are not, and the locations of the copper(A) tyrosinase mutations should be considered less precise. Two tyrosinase mutations in the copper(A) binding site of human tyrosinase are within conserved amino acid motifs found in most tyrosinases and hemocyanin copper binding sites (Müller et al., 1988). The mutation A206T is found in the motif Pro-X-Phe-X-X-X-His, between the proline and the phenylalanine, causing a change from an alanine to a threonine, and the mutation F176I is within the motif Phe-X-X-XHis, changing the highly conserved phenylalanine to an isoleucine. Both these motifs are thought to be involved in the correct placement of the histidine ligands responsible for copper binding. The other mutations are found to be either within the a-helix or at the junction of the a-helix and the connecting loop structure. Our current hypothesis is that these mutations disrupt correct copper atom orientation by displacing the a-helices relative to each other (Oetting and King, 1992b). Spritz et al. (1997) reported that some missense mutations disrupted copper binding, resulting in an inactive protein. There were two missense mutations, P406L and R402Q, that did exhibit copper binding but still had no detectable enzymatic activity. The action of these mutations 223

CHAPTER 11

may either prevent one of the copper atoms from binding or displace the copper atoms by altering the copper to copper distance. Either case would prevent the formation of oxytyrosinase and render the enzyme inactive. The limited number of mutations identiﬁed to date in TRP-1 and TRP-2 preclude a similar prediction of structure–function relationship for these proteins. There is also evidence that tyrosinase mutations result in retention of the enzyme in the endoplasmic reticulum and subsequent degradation, leading to speculation that albinism is, in part, an ER retention disease (Halaban et al., 1997, 2000; Toyofuku et al., 2001a, b).

Perspectives The tyrosinase gene family provides an excellent model for understanding a number of biological phenomena. These include gene evolution, the biology of a complex biochemical pathway, protein sorting, and protein–protein interaction. Two of these genes, TYR and TRP1, are involved in albinism. Analysis of tyrosinase mutations associated with oculocutaneous albinism type I has provided insight into the spectrum of phenotypes associated with a single gene disorder.

Acknowledgments V. S. acknowledges the support of NIH grant R01 AR048913.

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Oetting, W. S., C. J. Witkop, S. A. Brown, R. Colomer, J. P. Fryer, K. E. Bloom, and R. A. King. A frequent mutation in the tyrosinase gene associated with type I-A (tyrosinase-negative) oculocutaneous albinism in Puerto Rico. Am. J. Hum. Genet. 52:17–23, 1993b. Oetting, W. S., J. P. Fryer, Y. Oofuji, L. R. Middendorf, J. A. Brumbaugh, C. G. Summers, and R. A. King. Analysis of tyrosinase gene mutations using direct automated infrared ﬂuorescence DNA sequencing of ampliﬁed exons. Electrophoresis 15:159–164, 1994. Oetting, W. S., W. Skach, J. P. Fryer, and R. A. King. Unusual tyrosinase mutations associated with OCA1. Am. J. Hum. Genet. 53:935, 1995. Orlow, S. J. Melanosomes are specialized members of the lysosomal lineage of organelles. J. Invest. Dermatol. 105:3–7, 1995. Orlow, S. J., M. L. Lamoreux, S. Pifko-Hirst, and B. K. Zhou. Pathogenesis of the platinum (cp) mutation, a model for oculocutaneous albinism. J. Invest. Dermatol. 101:137–140, 1993a. Orlow, S. J., R. E. Boissy, D. J. Moran, and S. Pifko-Hirst. Subcellular distribution of tyrosinase and tyrosinase-related protein-1: implications for melanosomal biogenesis. J. Invest. Dermatol. 100:55–64, 1993b. Orlow, S. J., B.-K. Zhou, A. K. Chakraborty, M. Drucker, S. PifkoHirst, and J. M. Pawelek. High-molecular-weight forms of tyrosinase and the tyrosinase-related proteins: evidence for a melanogenic complex. J. Invest. Dermatol. 103:196–201, 1994. Pawelek, J. M. After DOPAchrome? Pigment Cell Res. 4:53–62, 1991. Pawelek, J. M., A. M. Körner, A. Bergstrom, and J. Bolognia. New regulators of melanin biosynthesis and the autodestruction of melanoma cells. Nature 286:617–619, 1980. Peng, G., J. D. Taylor, and T. T. Tchen. Goldﬁsh tyrosinase related protein I (TRP-1): Deduced amino acid sequence from cDNA and comments on structural features. Pigment Cell Res. 7:9–16, 1995. Ponnazhagan, S., L. Hou, and B. S. Kwon. Structural organization of the human tyrosinase gene and sequence analysis and characterization of its promoter region. J. Invest. Dermatol. 102:744–748, 1994. Porter, S. D., and C. J. Meyer. A distal tyrosinase upstream element stimulates gene expression in neural-crest-derived melanocytes of transgenic mice: position-independent and mosaic expression. Development 120:2103–2111, 1994. Porter, S., L. Larue, and B. Mintz. Mosaicism of tyrosinase-locus transcription and chromatin structure in dark vs light melanocyte clones of homozygous chinchilla-mottled mice. Dev. Genet. 12:393–402, 1991. Pruitt, K. D., and D. R. Maglott. RefSeq and LocusLink: NCBI genecentered resources. Nucleic Acids Res. 29:137–140, 2001. Regales, L., P. Giraldo, A. Garcia-Diaz, A. Lavado, and L. Montoliu. Identiﬁcation and functional validation of a 5¢ upstream regulatory sequence in the human tyrosinase gene homologous to the locus control region of the mouse tyrosinase gene. Pigment Cell Res. 16:685–693, 2003. Rosenberg, S. A. Progress in human tumor immunology and immunotherapy. Nature 411:380–384, 2001. Sato, S., H. Miura, H. Yamamoto, and T. Takeuchi. Identiﬁcation of nuclear factors that bind to the mouse tyrosinase gene regulatory region. Pigment Cell Res. 7:279–284, 1994. Schedl, A., L. Montollu, G. Kelsey, and G. Schütz. A yeast artiﬁcial chromosome covering the tyrosinase gene confers copy numberdependent expression in transgenic mice. Nature 362:258–261, 1993. Schmidt, A., and F. Beermann. Molecular basis of dark-eyed albinism in the mouse. Proc. Natl. Acad. Sci. USA 91:4756–4760, 1994. Setaluri, V. Sorting and targeting of melanosomal membrane proteins: signals, pathways and mechanisms. Pigment Cell Res. 113:128– 134, 2000.

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CHAPTER 11 Shibahara, S., Y. Tomita, T. Sakakura, C. Nager, B. Chaudhuri, and R. Muller. Cloning and expression of cDNA encoding mouse tyrosinase. Nucleic Acids Res. 14:2413–2427, 1986. Shibahara, S., S. Okinaga, Y. Tomita, A. Takeda, H. Yamamoto, M. Sato, and T. Takeuchi. A point mutation in the tyrosinase gene of BALB/c albino mouse causing the cysteine to serine substitution at position 85. Eur. J. Biochem. 189:455–461, 1990. Shibahara, S., H. Taguchi, R. M. Muller, K. Shibata, T. Cohen, Y. Tomita, and H. Tagami. Structural organization of the pigment cellspeciﬁc gene located at the brown locus in mouse. Its promoter activity and alternatively spliced transcript. J. Biol. Chem. 266: 15895–15901, 1991. Shibata, K., Y. Muraosa, Y. Tomita, H. Tagami, and S. Shibahara. Identiﬁcation of a cis-acting element that enhances the pigment cellspeciﬁc expression of the human tyrosinase gene. J. Biol. Chem. 267:20584–20588, 1992a. Shibata, K., K. Takeda, Y. Tomita, H. Tagami, and S. Shibahara. Downstream region of the human tyrosinase-related protein gene enhances its promoter activity. Biochem. Biophys. Res. Commun. 184:568–575, 1992b. Silvers, W. K. The Coat Colors of Mice. A Model for Mammalian Gene Action and Interaction. New York: Springer-Verlag, 1979. Simmen, T., A. Schmidt, W. Hunzilker, and F. Beermann. 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, 1999. Solano, F., J. H. Martinez-Liarte, C. Jimenez-Cervantes, J. C. GarciaBorron, and J. A. Lozano. Dopachrome tautomerase is a zinccontaining enzyme. Biochem. Biophys. Res. Commun. 204:1243– 1250, 1994. Spritz, R. A., L. Ho, M. Furumura, and V. J. Hearing. Mutational analysis of copper binding by human tyrosinase. J. Invest. Dermatol. 109:207–212, 1997. Steel, K. P., D. R. Davidson, and I. J. Jackson. TRP2/DT, a new early melanoblast marker, shows that steel growth factor (c-kit ligand) is a survival factor. Development 115:1111–1119, 1992. Sturm, R. A., B. J. O’Sullivan, J. A. F. Thomson, N. Jamshidi, J. Pedley, and P. Parsons. Expression studies of pigmentation and POUdomain genes in human melanoma cells. Pigment Cell Res. 7:235–240, 1994a. Sturm, R. A., E. Baker, and G. R. Sutherland. Assignment of the tyrosinase-related protein-2 gene (TYRP2) to human chromosome 13q31–q32 by ﬂuorescence in situ hybridization: extended synteny with mouse chromosome 14. Genomics 21:293–296, 1994b. Sturm, R. A., B. J. O’Sullivan, N. F. Box, A. G. Smith, S. E. Smit, E. R. J. Puttick, P. G. Parsons, and I. S. Dunn. Chromosomal structure of the human TYRP1 and TYRP2 loci and comparison of the tyrosinase-related protein gene family. Genomics 29:24–34, 1995. Takeda, A., J. Matsunaga, Y. Tomita, H. Tagami, and S. Shibahara. Molecular analysis of the DNA segments cross-hybridizable to the tyrosinase gene in patients afﬂicted with oculocutaneous albinism. Tohoku J. Exp. Med. 159:333–340, 1989a. Takeda, A., Y. Tomita, S. Okinaga, H. Tagami, and S. Shibahara. Functional analysis of the cDNA encoding human tyrosinase precursor. Biochem. Biophys. Res. Commun. 162:984–990, 1989b. Takeda, A., J. Matsunaga, Y. Tomita, H. Tagami, and S. Shibahara. Nucleotide sequence of the putative human tyrosinase pseudogene. Tohoku J. Exp. Med. 163:295–297, 1991. Takeuchi, H., C. Kuo, D. L. Morton, H. J. Wang, and D. S. Hoon. Expression of differentiation melanoma-associated antigen genes is associated with favorable disease outcome in advanced-stage melanomas. Cancer Res. 63:441–448, 2003. Tamate, H. B., T. Hirobe, K. Wakamatsu, S. Ito, S. Shibahara, and K. Ishikawa. Levels of tyrosinase and its mRNA in coat-color mutants of C57BL/10J congenic mice: effects of genic substitution at the agouti, brown, albino, dilute and pink-eyed dilution loci. J. Exp. Zool. 250:304–311, 1989.

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Taniguchi, M., T. Iwamoto, I. Nakashima, A. Nakayama, O. Masaharu, M. Matsuyama, and M. Takahashi. Establishment and characterization of a malignant melanocytic tumor cell line expressing the ret oncogene. Oncogene 7:1491–1496, 1992. Tassabehji, M., V. E. Newton, and A. P. Read. Waardenburg syndrome type 2 caused by mutations in the human microphthalmia (MITF) gene. Nature Genet. 8:251–255, 1994. Tomita, Y., A. Takeda, S. Okinaga, H. Tagami, and S. Shibahara. Human oculocutaneous albinism caused by a single base insertion in the tyrosinase gene. Biochem. Biophys. Res. Commun. 164:990– 996, 1989. Toyofuku, K., I. Wada, J. C. Valencia, T. Kushimoto, V. J. Ferrans, and V. J. Hearing. Oculocutaneous albinism type 1 and 3 are ER retention diseases: mutation of tyrosinase or Tyrp1 can affect the processing of both mutant and wildtype proteins. FASEB J. 15:2149–2161, 2001a. Toyofuku, K., I. Wada, R. A. Spritz, and V. J. Hearing. 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, 2001b. Tripathi, R. K., V. J. Hearing, K. Urabe, P. Aroca, and R. A. Spritz. Mutational mapping of the catalytic activities of human tyrosinase. J. Biol. Chem. 267:23707–23712, 1992a. Tripathi, R. K., K. M. Strunk, L. B. Giebel, R. G. Weleber, and R. A. Spritz. Tyrosinase gene mutations in Type I (tyrosinase-deﬁcient) oculocutaneous albinism deﬁne two clusters of missense substitutions. Am. J. Med. Genet. 43:865–871, 1992b. Tsukamoto, K., I. J. Jackson, K. Urabe, P. M. Montague, and V. J. Hearing. A second tyrosinase-related protein, TRP2, is a melanogenic enzyme termed DOPAchrome tautomerase. EMBO J. 11:519–526, 1992a. Tsukamoto, K., M. Jiménez, and V. J. Hearing. The nature of tyrosinase isozymes. In: The Pigment Cell: From the Molecular to the Clinical Level, Y. Mishima (ed.). Copenhagen: Munksgaard Press, 1992b, pp. 84–89. Urquhart, A. Human tyrosinase-like protein (TYRL) carboxy terminus: closer homology with the mouse protein than previously reported. Nucleic Acids Res. 19:5803, 1993. Vijayasaradhi,S., B. Bouchard, and A. N. Houghton. The melanoma antigen gp75 is the human homologue of the mouse b (brown) locus gene product. J. Exp. Med. 171:1375–1380, 1990. Vijayasaradhi, S., Y. X. Brigitte, B. Bouchard, and A. N. Houghton. Intracellular sorting and targeting of melanosomal membrane proteins: Identiﬁcation of signals for sorting of the human brown locus protein, GP75. J. Cell Biol. 130:807–820,1995a. Vijayasaradhi, S., P. M. Doskoch, J. Wolchok, and A. N. Houghton. Melanocyte differentiation marker gp75, the brown locus protein, can be regulated independently of tyrosinase and pigmentation. J. Invest. Dermatol. 105:113–119, 1995b. Wang, R. F., P. F. Robbins, Y. Kawakami, X. Q. Kang, and S. A. Rosenberg. Identiﬁcation of a gene encoding a melanoma tumor antigen recognized by HLA-A31-restricted tumor-inﬁltrating lymphocytes. J. Exp. Med. 181:799–804, 1995. Wang, R. F., E. Appella, Y. Kawakami, X. Kang, and S. A. Rosenberg. Identiﬁcation of TRP-2 as a human tumor antigen recognized by cytotoxic T lymphocytes. J. Exp. Med. 184:2207–2216, 1996. Wilcox, D. E., A. G. Porras, Y. T. Hwang, K. Lerch, M. E. Winkler, and E. I. Solomon. Substrate analogue binding to the coupled binuclear copper active site in tyrosinase. J. Am. Chem. Soc. 107:4015–4027, 1995. Winder, A. J., T. Kobayashi, K. Tsukamoto, K. Urabe, P. Aroca, K. Kameyama, and V. J. Hearing. The tyrosinase gene family: Interactions of melanogenic proteins to regulate melanogenesis. Cell. Mol. Biol. Res. 40:613–626, 1994a. Winder, A. J., A. Wittbjer, G. Odh, E. Rosengren, and H. Rorsman. The mouse brown (b) locus protein functions as a dopachrome tautomerase. Pigment Cell Res. 7:305–310, 1994b.

THE TYROSINASE GENE FAMILY Wittbjer, A., B. Dahlback, G. Odh, A. M. Rosengren, E. Rosengren, and H. Rorsman. Isolation of human tyrosinase from cultured melanoma cells. Acta Derm. Venereol. Suppl. 69:125–131, 1989. Yamamoto, H., S. Takeuchi, T. Kudo, K. Makino, A. Nakata, T. Shinoda, and T. Takeuchi. Cloning and sequencing of mouse tyrosinase cDNA. Jpn. J. Genet. 62:271–274, 1987. Yamamoto, H., T. Kudo, N. Masuko, H. Miura, S. Sato, M. Tanaka, S. Tanaka, S. Takeuchi, S. Shibahara, and T. Takeuchi. Phylogeny of regulatory regions of vertebrate tyrosinase genes. Pigment Cell Res. 5:284–294, 1992. Yasumoto, K.-I., K. Yokoyama, K. Shibata, Y. Tomita, and S. Shibahara. Microphthalmia-associated transcription factor as a regulator for melanocyte-speciﬁc transcription of the human tyrosinase gene. Mol. Cell. Biol. 14:8058–8070, 1994. Yasumoto, K.-I., H. Mahalingam, H. Suzuki, M. Yoshizawa, K. Yokoyama, and S. Shibahara. Transcriptional activation of the melanocyte-speciﬁc genes by the human homolog of

the mouse microphthalmia protein. J. Biochem. 118:874–881, 1995. Yokoyama, T., D. W. Silversides, K. G. Waymire, B. S. Kwon, T. Takeuchi, and P. A. Overbeek. Conserved cysteine to serine mutation in tyrosinase is responsible for the classical albino mutation in laboratory mice. Nucleic Acids Res. 18:7293–7298, 1990. Yokoyama, K., H. Suzuki, K.-I. Yasumoto, Y. Tomita, and S. Shibahara. Molecular cloning and functional analysis of a cDNA coding for human DOPAchrome tautomerase/tyrosinase-related protein-2. Biochim. Biophys. Acta 1217:317–321, 1994. Zdarsky, E., J. Favor, and I. J. Jackson. The molecular basis of brown, an old mouse mutation, and of an induced revertant to wild-type. Genetics 126:443–449, 1990. Zhou, B. K., R. E. Boissy, S. Pifko-Hirst, D. J. Moran, and S. J. Orlow. Lysosome-associated membrane protein-1 (LAMP-1) is the melanocyte vesicular membrane glycoprotein band II. J. Invest. Dermatol. 100:110–114, 1993.

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12

Molecular Regulation of Melanin Formation: Melanosome Transporter Proteins Murray H. Brilliant

Summary 1 To make melanin, tyrosinase must be correctly targeted to the melanosome, its substrate, tyrosine, must be transported into the melanosome, and conditions that favor the enzymatic activity of tyrosinase must exist inside the melanosome. All these processes are affected by pH and ionic conditions. Therefore, ion transport across the melanosome membrane is critical for the biosynthesis of melanin. Two proteins, the P protein [encoded by the mouse pink-eyed dilution (p) locus and the human P locus] and MATP (membrane-associated transport protein, previously known as AIM-1, encoded by the formerly named mouse underwhite locus) are critical for normal pigmentation and are likely to be involved in ion ﬂux across the melanosomal membrane. 2 The mouse pink-eyed dilution locus, p, and its human homolog, P, encode a protein of 110 kDa with 12 membranespanning domains that is localized to the melanosome membrane, although a signiﬁcant fraction is thought to be associated with the endoplasmic reticulum. The p protein is required for normal melanin (primarily eumelanin) biosynthesis. Although the exact function of the p protein is currently unknown, it has a predicted structure in common with anion transporters and Na+/H+ antiporters. Melanocytes lacking a functional p protein have immature melanosomes with aberrant pH that only accumulate minute amounts of primarily eumelanin. In these p mutant melanocytes, tyrosinase is incorrectly processed and targeted. 3 Mutations of the human P gene on chromosome 15q11.2–q12 lead to tyrosinase-positive oculocutaneous albinism, deﬁned as OCA2, a recessive genetic disorder present in the general population at 1:38 000. OCA2 is more common among individuals of African origin, where a predominant haplotype is associated with a 2.7-kb deletion that includes exon 7 of the P gene. In some African populations, this deletion allele comprises up to 80% of the P gene mutations. A larger deletion of different origin is commonly associated with OCA2 among the Navajo people, an independent population that exhibits a high frequency of albinism. 4 The only known function of the P gene is in pigmentation. However, deletions of the human chromosomal interval 15q11q13 that contains the P gene lead to distinct phenotypes known as Prader–Willi syndrome or Angelman syndrome, 230

depending on whether the deletion is paternal or maternal in origin respectively. Although the critical regions and/or genes associated with these imprinting syndromes are distinct from each other and from the P gene, one homolog of the P gene is often deleted in PWS and AS patients. Those AS and PWS patients who are hemizygous for the P gene (in the context of a large deletion of 15q) often exhibit hypopigmentation relative to other family members, a puzzling outcome as carriers of null alleles of the P gene (in the context of a normal chromosome 15) are normally pigmented. Many radiation-induced alleles of the mouse p locus on chromosome 7 are associated with nonpigmentation phenotypes. All these other phenotypes are associated with deletions extending into neighboring genes. 5 The Matp (membrane-associated transporter protein) gene encodes a protein of ~58 kDa with 12 membrane-spanning domains that is very likely associated with the melanosome membrane. The encoded protein, Matp, likely functions as a transporter. One of its closest orthologs encodes a plant sucrose/proton transporter, and the Matp protein may play a role in pH and/or osmotic regulation of the melanosome. In Matp mutant melanocytes (as in p mutant melanocytes), tyrosinase is incorrectly processed and targeted. Matp previously shared the name of AIM-1 with at least two other distinct and unrelated proteins. The mouse locus encoding Matp was formerly called the underwhite (uw) locus. 6 Various mouse mutant alleles of Matp (underwhite) lead to a spectrum of hypopigmented phenotypes, most of which are recessive, but one (MatpUw-dbr) is semi-dominant. 7 Mutations in the human MATP gene are associated with a form of oculocutaneous albinism, deﬁned as OCA4. While rare in the worldwide population, OCA4 is more common in the Japanese population, and cases have been described in the Turkish and German populations. 8 Variations in both the MATP and the P genes are associated with normal variations in human pigmentation.

Historical Background Mouse coat color genes have played an important role in understanding basic aspects of mammalian genetics. For example, the original p mutation and a mutation of the tyrosinase gene (c locus) were used to deﬁne the ﬁrst genetic linkage

MOLECULAR REGULATION OF MELANIN FORMATION: MELANOSOME TRANSPORTER PROTEINS

group in the mouse (Haldane et al., 1915), now assigned to mouse chromosome 7 (Brilliant et al., 1996). The p locus was also used as a marker in transplantation studies to deﬁne histocompatability antigens (Snell and Stevens, 1961). The original Matp mutant allele, Matpuw, was used as a coat color marker in many gene linkage and mapping studies (Davisson et al., 1990). The existence of genes that are associated with tyrosinasepositive albinism was hypothesized long ago (Witkop et al., 1970). The P and MATP genes are now known to be the most common genes mutated in tyrosinase-positive albinism, causing OCA2 (Gardner et al., 1992; Rinchik et al., 1993) and OCA4 (Newton et al., 2001) respectively.

Current Concepts Overview Melanin biosynthesis occurs in melanosomes, thought to be derived from the fusion of at least two vesicle compartments, including endosome-derived vesicles that share some features with lysosomes (Dell’Angelica et al., 2000; Orlow, 1995). The initial and rate-limiting step in melanin biosynthesis is catalyzed by tyrosinase, converting tyrosine to DOPAquinone (Cooksey et al., 1997; Hearing and Tsukamoto, 1991). Mature (highly pigmented) “Stage IV” melanosomes derive from earlier stages. The ﬁrst recognizable stage is the premelanosome, which is distinguished by a disorganized luminal matrix and no melanin. Stage I melanosomes also lack melanin, but are deﬁned by a parallel array of the luminal matrix. Stages II and III are characterized by increased melanin deposition, and Stage IV melanosomes contain the most melanin. Although mouse skin melanosomes are acidic (Bhatnagar et al., 1993; Devi et al., 1987; Ramaiah, 1996), the luminal pH is probably not static as melanosomes develop, and differences in melanosome pH in human melanocytes have been reported between those of Caucasian and black origin (Fuller et al., 2001; Smith et al., 2004). To make melanin: (1) tyrosinase must be correctly processed; (2) tyrosine must be targeted to the melanosome; and (3) conditions that favor the enzymatic activity of tyrosinase must exist inside the melanosome. All these processes are affected by pH (Ancans and Thody, 2000; Ancans et al., 2001; Halaban et al., 2002; Manga and Orlow, 2001; Potterf et al., 1998; Ramaiah, 1996; Watabe et al., 2004). The product of the p (pink-eyed dilution) gene is likely an ion transporter (Gardner et al., 1992; Rinchik et al., 1993) that plays a critical role in melanin biosynthesis through the regulation of melanosome pH (Puri et al., 2000). Matp (membraneassociated transport protein, encoded by the mouse locus previously known as underwhite, uw) also plays a key role in melanin biosynthesis (Lehman et al., 2000; Sweet et al., 1998). Matp likely mediates the transport of an unknown, but critical substance across the melanosome membrane and has high homology at the amino acid level to plant proton/ sucrose transporters (Newton et al., 2001).

The Mouse Pink-eyed Dilution (p) Gene and Its Alleles The original mutant p allele probably arose in Manchuria or Japan in Mus molosinus (Brilliant et al., 1994a) and was incorporated into several common laboratory strains of mice (e.g. SJL/J, 129/J, P/J, and FS/Ei) during the early twentieth century. The ﬁrst mutant allele of the p locus to be characterized molecularly was the pun (pink-eyed unstable) allele, which exhibits high-frequency (somatic) reversion to wild type (Melvold, 1971). Using the technique of genome scanning, the pun allele was found to result from a duplication of 75 kb of genomic DNA (Brilliant et al., 1991; Gondo et al., 1993) that includes exons 6–18 (Oakey et al., 1996) of the 24-exon p gene. A fragment of genomic DNA that cross-hybridized to various mammalian species (now know to include exon 19) was used to screen a melanoma cDNA library to obtain the p gene cDNA and its human homolog (Gardner et al., 1992). Parallel studies using a candidate gene approach found that a previously unknown human cDNA fragment (DN10) identiﬁed an alternative exon of the human P gene (Lee et al., 1995; Rinchik et al., 1993). Many mouse mutant p alleles have been characterized at the molecular level (Culiat et al., 1993, 1994; Gardner et al., 1992; Johnson et al., 1995; Nakatsu et al., 1993; Oakey et al., 1996; Rinchik et al., 1993). The discovery of the human P cDNA has led to an understanding of the molecular basis for tyrosinase-positive oculocutaneous albinism (OCA2). More than 100 p alleles have been identiﬁed, some of which were de novo in origin and some of which were induced by X-rays or chemical mutagens (Lyon et al., 1992; Russell et al., 1995; M. F. Lyon, personal communication). In the homozygous state, each mutant p allele causes hypopigmentation ranging from a minor reduction in coat color to a dramatic reduction in both coat and eye color characteristic of the original p mutation. In addition to affecting pigmentation, several mutant alleles are associated with other abnormalities, including decreased neonatal viability, neurologic disorders, cleft palate, male sterility, female semi-fertility, viability, and prenatal lethality (Brilliant, 1992; Culiat et al., 1993, 1994; Hagiwara et al., 2000; Lyon et al., 1992; Nakatsu et al., 1993; Russell et al., 1995). All the mutations with these additional phenotypes were induced by radiation, and all affect surrounding genes (Johnson et al., 1995; Lyon et al., 1992; Russell et al., 1995) or are the result of a chromosomal inversion (Hagiwara et al., 2000). Mutations of the p gene alone cause effects on pigmentation only (Gardner et al., 1992; Johnson et al., 1995; Lyon et al., 1992; Russell et al., 1995). The p mutations result in a dramatic reduction in melanin (primarily eumelanin). In the hair, p pigment granules have been described as shred-like (Russell, 1949). The same description has been applied to the small irregular shaped melanosomes in the harderian gland of p mice (Markert and Silvers, 1956). In p mice, premelanosomes from embryonic choroid and retina are ﬁbrillar in appearance, as are those 231

CHAPTER 12

CHOROID

RPE

CHOROID

RPE

Fig. 12.1. Electron micrograph of wild-type and p/p mutant eyes. A wild-type eye is shown on the left; a p/p mutant eye is shown on the right. Note the aberrant sizes and minimal pigment content of the melanosomes (arrows) in the choroid and retinal epithelium (RPE) of the p/p eye, a feature shared with other forms of albinism.

from the adult choroid. However, most melanosomes from the adult retina are more particulate in appearance. Most p melanosomes show some pigment, indicated as an increase in electron density and diameter of melanoﬁlaments, but none is fully melanized (Hearing et al., 1973). The melanosomes within p/p melanocytes appear as Stage 1 and Stage 2 melanosomes, as described in the above references (Fig. 12.1). Transplantation studies in the mouse have demonstrated that the p defect is intrinsic to melanocytes (Stephenson and Hornby, 1985).

The p Gene Protein and Its Function Molecular analysis of the p transcript and protein have corroborated much of the past phenotypic data and have extended our knowledge about the p protein’s role in pigmentation, even if its precise function remains unknown at this time. Northern blot analyses have conﬁrmed that melanocytes are the predominant p gene-expressing cell type, with lowlevel expression of the p gene in brain, testes, and ovaries (Gardner et al., 1992; Rinchik et al., 1993). The expression of the p gene in testes and ovaries is also conserved in medaka ﬁsh (Fukamachi et al., 2004). However, the signiﬁcance of p gene expression in nonpigmented tissues is unknown as animals and humans lacking P gene expression are normally fertile, and any neurologic problems are limited to the visual system, the same problems as those commonly associated with other forms of albinism. The size of the mouse p gene transcript is 3.3 kb, encoding a predicted protein of 833 amino acids (Gardner et al., 1992; Rinchik et al., 1993). The mouse p gene product, like its 232

human homolog P, encodes a protein with 12 membranespanning domains. Using antisera against a synthetic peptide from the ﬁrst luminal loop (amino acids 285–298), Rosemblat et al. (1994) were able to characterize the p protein as an integral, melanosomal membrane protein of 110 kDa that does not appear to be glycosylated (tunicamycin treatment does not alter its gel mobility). Subsequent studies (Chen et al., 2002) suggested that a signiﬁcant fraction of the p protein is associated with the endoplasmic reticulum (ER). Whether this fraction of the p protein functions in the ER or is in transit is unknown. The p protein may be speciﬁcally targeted to the melanosome or one of its precursors. In normal pigmented melanocytes, a fraction of the p protein is present in an intracellular compartment distinct from that containing tyrosinase and Trp-1. It has also been noted that melanosomes lacking p protein are lacking a high-molecular-weight complex of the p protein, tyrosinase, and Trp-1, and possess characteristics of immature premelanosomes (Rosemblat et al., 1994). Thus, in addition to a potential transport function, the p protein may also play a critical role in the biogenesis of normal melanosomes, perhaps by providing conditions (e.g. pH) favorable for their proper structure and maturation. The melanosome, as a specialized endosome, may derive its protein components by means of signals and sorting mechanisms that distinguish it from a lysosome. In fact, a speciﬁc protein sequence motif, capable of targeting Trp-1 to the melanosomal membrane, exists within its carboxyl-terminus (Vijayasaradhi et al., 1995). When this Trp-1 amino acid sequence was incorporated in chimeric proteins, they were correctly targeted. The sequence capable of targeting Trp-1 is conserved across species, and a related sequence is found in several other melanosome proteins, perhaps including the p protein. However, it remains to be shown whether the sequence in the p protein related to the Trp-1 sorting sequence actually functions in the same way. If so, it would confound the observation that a subset of the p protein is in a different intracellular compartment from tyrosinase and Trp-1 (Chen et al., 2002; Rosemblat et al., 1994). The p protein may interact with melanin. The p protein (along with the silver protein) is far less extractable from melanized melanosomes (in melanocytes or in a cell-free assay system) than it is from poorly melanized melanosomes (Donatien and Orlow, 1995). It may be that some of the loops of the p protein that protrude inside the melanosome are somehow trapped within the melanin polymer. If this close association with melanin impedes p gene function, then perhaps this is one way to limit the melanin content of the developing melanosome. An interesting observation is that the p transcript is expressed in the black dorsal skin, but not the yellow ventral skin in agouti at/at mice (Rinchik et al., 1993), corroborating earlier notions that the p mutations affect eumelanin but not pheomelanin biosynthesis. Eumelanin biosynthesis is favored when the melanocortin 1 receptor (Mc1r) is stimulated by aMSH and upregulates cAMP; pheomelanin is favored when

MOLECULAR REGULATION OF MELANIN FORMATION: MELANOSOME TRANSPORTER PROTEINS

this signaling is antagonized by the agouti protein. Expression of the p gene is upregulated by a-MSH and db-cAMP (a synthetic analog of cAMP), implying that the p protein plays a major role in the eumelanin/pheomelanin switch (Ancans et al., 2003). From its predicted protein structure, the p protein is likely to be a transporter critical to melanocyte function (Gardner et al., 1992; Rinchik et al., 1993). The p protein was proposed to be a tyrosine transporter (Rinchik et al., 1993), based on sequence homology that has subsequently been shown not to be signiﬁcant (Lee et al., 1995; Rosemblat et al., 1994). Supporting the hypothesis, however, are the observations that both mouse and human melanocytes can be induced to pigment in vitro with high tyrosine (Sidman and Pearlstein, 1965; Witkop et al., 1973). This observation was also conﬁrmed using cultured mouse skin melanocytes (Potterf et al., 1998; Rosemblat et al., 1998). It is possible that the increase in pigmentation detected in mutant p cells when exposed to very high concentrations of tyrosine results from driving the reaction by substantially increasing the substrate. Moreover, the melanin produced in these cells does not appear to be synthesized in melanosomes, and may result from the activity of mislocalized tyrosinase (Manga and Orlow, 1999; Potterf et al., 1998). Most importantly, direct biochemical assays show no difference in tyrosine transport between normal and p melanocytes at the level of the melanocyte or the melanosome. Plasma membrane tyrosine transport was found to be normal in p (pcp/pcp) melanocytes (Km 89 mM; Vmax 302 pmol/min/mg cell protein), and the melanosome-rich granular fractions of normal (melan-a) and p melanocytes (pcp/pcp) were essentially the same, taking up 10 mM [3H] tyrosine at about 21 pmol/min/mg protein (Gahl et al., 1995). Thus, although the p protein may be a transporter, it does not contribute signiﬁcantly to tyrosine transport. The most homologous bacterial proteins to p include the Staphylococcus aureus and Escherichia coli ArsB proteins (the anion-conducting pathway of a group of proteins that together confer resistance to arsenic), the E. coli Na+/H+ antiporter, and protein 38L from Mycobacterium leprae (Lee et al., 1995; Rosemblat et al., 1994). It may be that the p protein is similarly involved in ion transport, potentially modulating the pH of the melanosome. Indeed, it was found that the mouse p gene can function in yeast and affect cellular sensitivity to arsenic and other metalloids and modulate intracellular glutathione metabolism (Staleva et al., 2002). Although distinct, melanosomes and lysosomes share at least partial endosomal origins, and both types of organelles are characterized by an acidic lumen (Dell’Angelica et al., 2000; Orlow, 1995). Mouse melanosomes have been reported to be very acidic, i.e. pH 3.5 (Bhatnagar et al., 1993; Devi et al., 1987; Moellmann et al., 1988; Tripathi et al., 1988) and, as such, may more closely resemble skin melanosomes from human Caucasians, whereas melanosomes from more darkly pigmented individuals (i.e. black Africans) may be more neutral in pH (Fuller et al., 2001; Smith et al., 2004). Additionally, the pH of melanosomes may be dynamic, with

early stage melanosomes more acidic and later stage melanosomes more neutral in pH allowing for higher tyrosinase activity. In general, acidiﬁcation of various intracellular compartments is important for a number of processes including receptor-mediated endocytosis, receptor recycling, and membrane trafﬁcking within the cell (Dautry-Varsat et al., 1983). Acidiﬁcation appears to be required for normal melanosome biogenesis and protein trafﬁcking (reviewed by Brilliant, 2001; Raposo and Marks, 2002), whereas a more neutral pH results in higher tyrosinase activity (Ancans and Thody, 2000; Ancans et al., 2001; Fuller et al., 2001; Manga and Orlow, 2001; Smith et al., 2004). The identity of all the molecular components mediating the pH of endosome-derived organelles is not known. However, it is thought that an anion channel is essential for the acidiﬁcation of vacuole compartments by ATP-driven proton pumps present in endosomes, Golgi-derived vesicles, and lysosomes (Al-Awqati, 1995). Anion (Cl–, SO4=, or HCO3–) conductance provides the compensating charge balance to electrogenic proton transport. It remains to be determined whether or not there is a melanosome-speciﬁc, ATP-driven proton pump. Nevertheless, the melanosome has a unique membrane protein, p, that may be an anion transporter or a Na+/H+ antiporter that may play a role in the pH regulation of melanosomes (Puri et al., 2000). The regulation of pH in melanosomes is likely to be complex and to involve additional proteins, such as Matp (discussed below). Alternative hypotheses for the function of the p protein have been proposed. The p protein may function to transport sulfhydryl compounds out of the melanosome to permit the formation of eumelanin (Lamoreux et al., 1995). The p protein has also been hypothesized to affect the rate of an existing melanin-synthesizing enzyme system to achieve normal pigmentation (Coleman, 1962; Sidman and Pearlstein, 1965). Other observations suggest that the p, tyrosinase, and Trp-1 (tyrosinase-related protein-1) proteins exist in a highmolecular-weight complex that is not formed in p mutant melanocytes (Chiu et al., 1993; Lamoreux et al., 1995). More recently, it has been shown in several studies that, in the absence of a normal p protein, tyrosinase is mistargeted and accumulates in the ER (Manga and Orlow, 1999; Potterf et al., 1998). Other studies imply that a signiﬁcant portion of the p protein is found in the ER, and their authors have put forward the hypothesis that the P protein is involved in tyrosinase maturation and folding in the ER (Chen et al., 2002; Toyofuku et al., 2002), which may also involve glutathione (Staleva et al., 2002). Melanocytes have a speciﬁc factor that is critical for tyrosinase maturation; however, as p-null melanocytes also appear to have this factor, the p protein itself appears to be this factor (Francis et al., 2003). Moreover, the absence of the Matp protein has a similar phenotype (Costin et al., 2003), suggesting that both these proteins may play an indirect role in the folding and targeting of tyrosinase, perhaps by modulating pH (Halaban et al., 2002; Watabe et al., 2004). In any case, the p (and Matp) protein plays a critical coordinating role in melanogenesis. 233

CHAPTER 12

OCA2 and the Human P Gene The human ortholog (P) of the pink-eyed dilution gene has been identiﬁed and characterized. Mutations of the human P gene are responsible for type II oculocutaneous albinism (OCA2). In addition, polymorphic variation in the human P gene is responsible, in part, for variations in human pigmentation. Oculocutaneous albinism (OCA) is characterized by abnormally low amounts of melanin in the eyes and skin (see Chapter 31). Abnormally low amounts of melanin in the developing eye lead to abnormal routing of optic nerve ﬁbers, resulting in strabismus and loss of binocular vision. Other ocular effects of albinism include photophobia, nystagmus, and foveal hypoplasia with reduced visual acuity. The reduction in skin pigmentation in individuals with OCA is associated with an increased sensitivity to ultraviolet radiation and a predisposition to skin cancer. Before molecular diagnostics, only two major types of OCA were recognized, tyrosinaserelated OCA and tyrosinase-positive OCA. Mutations of the human tyrosinase gene on chromosome 11 lead to tyrosinaserelated albinism, deﬁned as OCA1 (Tomita et al., 1989; reviewed by King et al., 2003a), whereas tyrosinase-positive albinism is genetically complex (see Chapter 31). In most populations, the major cause of tyrosinase-positive albinism is mutation of the P gene (reviewed by Oetting et al., 1998; see Chapter 31), although mutations of the Matp gene cause a similar phenotype (Newton et al., 2001). The human P gene is on chromosome 15q in a region demonstrated to be linked to OCA2 in native South Africans (Kedda et al., 1994; Ramsay et al., 1992). Mutations in the human P gene lead to tyrosinase-positive OCA, deﬁned as OCA2 (Durham-Pierre et al., 1994; Lee et al., 1994a, b; Rinchik et al., 1993). The phenotype of OCA2 is broad, ranging from minimal to moderate pigmentation of the hair, skin, and iris. The skin pigment tends to be localized in freckles, lentigines, or nevi rather than generalized, and the ability to tan is not well deﬁned (Oetting and King, 1999; see Chapter 31). In contrast to OCA1, individuals with OCA2 usually have pigmented hair at birth that tends to darken somewhat with age. A subtype of albinism known as brown oculocutaneous albinism (BOCA), described in African populations, has been shown to be associated with heterozygosity of a partially functioning allele and a null allele (Kerr et al., 2000; Manga et al., 2001). Additionally, at least one case of autosomal recessive ocular albinism (AROA) is the result of P gene mutations (Lee et al., 1994a). Thus, the phenotypic range of P gene mutations is broad. Moreover, normal MC1R (melanocortin receptor 1) variants dramatically inﬂuence the OCA2 phenotype (King et al., 2003b). In the general population of the United States, tyrosinasepositive OCA occurs in 1:30 000 Caucasians and in 1:17 000 blacks (Witkop, 1985). P gene mutations (OCA2) have been detected in these and other racial groups. There are several

234

genetic isolates with a very high frequency of tyrosinasepositive OCA, e.g. the Brandywine, Maryland isolate (1:85) and several native North American Pueblo Indian groups: the Zuni, Hopi, and Jemez people (approximately 1:240; Witkop, 1985; Witkop et al., 1972; Woolf and Dukepoo, 1969), as well as the Navajo people (approximately 1:2000; Yi et al., 2003) and several native South American Indian groups (reviewed by Jeambrun and Sergent, 1991). Presumably, because these represent small restricted populations, individuals within these populations are homozygous for the same recessive mutation of a gene required for normal pigmentation. The P gene is a likely candidate for the OCA seen in all these groups. Indeed, individuals with tyrosinase-positive OCA from the Brandywine, Maryland isolate are homozygous for a deletion allele of the P gene (a 2.7-kb deletion that includes exon 7; DurhamPierre et al., 1994), and Navajos with OCA are typically homozygous for an even larger deletion (122.5 kb) of the P gene. However, it is formally possible that mutations in another gene (e.g. MATP) lead to a similar phenotype of tyrosinase-positive OCA in individuals and groups for which no molecular data are yet available. The tyrosinase-positive OCA phenotype in Africans and African-Americans is characterized by yellow hair, white skin (sometimes with localized pigmented ephelides; Stevens et al., 1995), and irises that are partially or completely pigmented (Oetting and King, 1999; see Chapter 31). This is the most common albinism phenotype because of the high frequency in these populations (ranging from 1:2000 to 1:5000 in large parts of subSaharan Africa). The most common mutation in this group (so far found exclusively in Africans and individuals of African ancestry) is a 2.7-kb deletion that removes exon 7 along with ﬂanking intron sequences, ﬁrst identiﬁed in the Brandywine, Maryland isolate (Durham-Pierre et al., 1994). It is estimated that this single mutation is associated with 25–50% of all mutant P alleles in African-Americans (Durham-Pierre et al., 1994, 1996), although other diverse mutant alleles have been described in this population (Lee et al., 1994b). The 2.7-kb deletion allele accounts for close to 80% of mutant P alleles in South Africa, Zimbabwe, Tanzania, and other parts of subSaharan Africa (Durham-Pierre et al., 1994; Lund et al., 1997; Puri et al., 1997; Spritz et al., 1995; Stevens et al., 1995, 1997). The phenotypic range of OCA2 is now being deﬁned through the molecular characterization of the gene in different individuals with albinism, and it is expected that P gene mutations in OCA2 will be diverse. The missense mutations described to date do not seem to cluster in any speciﬁc region of the peptide, as observed for tyrosinase, but most mutations described so far are in the carboxy half of the polypeptide that contains the majority of the 12 membrane-spanning domains (Oetting et al., 1998). A signiﬁcant portion of the P missense mutations is found at amino acids conserved between the mouse and human P genes and a group of bacterial transport proteins with 12 membrane-spanning domains (Lee et al., 1995). The size of the human P gene transcript is 3.4 kb, encoding

MOLECULAR REGULATION OF MELANIN FORMATION: MELANOSOME TRANSPORTER PROTEINS

a predicted protein of 838 amino acids (Rinchik et al., 1993). Both the human and the mouse proteins encode a 12 membrane-spanning domain protein of unknown function, but related to a group of transport proteins (Gardner et al., 1992; Lee et al., 1995; Rinchik et al., 1993; Rosemblat et al., 1994). The P gene is encoded by 24 exons (plus one alternate exon that contains an in-frame stop codon, corresponding to IR10-1, an anonymous genomic clone) that spans approximately 250 to 650 kb of genomic DNA (Lee et al., 1995). The human proximal promoter region contains sequences that might be binding sites for transcription factors including the following 12 motifs: one AP4, four discrete and one complex AP2, one CF1, one GCF, three SP1, and one TFIID. No TATA or CCAAT motifs or melanocyte-speciﬁc motifs (i.e. M-box) have been described (Lee et al., 1995).

In addition to observing hypopigmentation in PWS and AS patients, several cases of PWS and AS are associated with OCA2 (Creel et al., 1986; Fridman et al., 2003; Fryburg et al., 1991; Wallis and Beighton, 1989), with deletion of one homolog of the P gene in the context of PWS or AS and inheritance of a mutation on the other homolog (Brilliant et al., 1994b; Rinchik et al., 1993). Another pigmentation disorder, hypomelanosis of Ito (HI), is genetically complex, with a subset of patients being mosaic for 15q anomalies (reviewed by Pellegrino et al., 1995). These HI patients have hypopigmented whorls, streaks, or patches in addition to some phenotypic similarities to PWS or AS. Just as in PWS and AS, the P gene is likely to underlie at least part of the hypopigmentation phenotype in those HI patients with 15q anomalies.

The P Gene in Prader–Willi Syndrome, Angelman Syndrome, Hypomelanosis of Ito, and Hypopigmentation.

Evolution and the p Gene

Prader–Willi syndrome (PWS) and Angelman syndrome (AS) are genetic diseases associated with chromosome 15q aberrations. PWS and AS both localize to the same chromosomal region, 15q11q13, although the critical regions for the two syndromes are distinct (reviewed by Knoll et al., 1993; Nicholls, 1993). Two common types of deletions are seen in PWS and AS patients (Christian et al., 1995), and both types disrupt the marker D15S12 (IR10) within the human P gene. PWS involves the loss of a paternal component of 15q11q13, with or without maternal disomy for 15q. The opposite inheritance pattern is seen in AS, as it is associated with a deletion of the maternal component of 15q11q13, with or without paternal disomy for 15q. Among the common clinical features of PWS are neonatal hypotonia and failure to thrive, hyperphagia leading to obesity (starting at 1–2 years), mental retardation, behavior problems, craniofacial abnormalities, and hypogonadism (especially in males) leading to infertility (Bray et al., 1983). The clinical features of AS include severe mental retardation, microcephaly, seizures, hypotonia, ataxia, and craniofacial abnormalities (Clayton-Smith, 1993; Magenis et al., 1990). Both syndromes are also associated with hypopigmentation, as many of these patients have much lighter skin, eye, and hair color than other family members (Butler, 1989; ClaytonSmith, 1993; Hittner et al., 1982; King et al., 1993; Saitoh et al., 1994; Weisner et al., 1987). The hypopigmentation is observed in most PWS and AS patients with deletions of 15q11q13, and more speciﬁcally among those with a disruption of D15S12, identiﬁed by the IR10 probe (Hamabe et al., 1991; Spritz et al., 1997). Thus, patients who are hemizygous for the P gene in the context of a large deletion of 15q are hypopigmented. This observation is difﬁcult to resolve with the recessive nature of both human P and mouse p mutations. Perhaps other genetic determinants of pigmentation are in the 15q11q13 interval. In contrast, people with extra copies of the P gene are hyperpigmented (Akahoshi et al., 2001, 2004).

There is substantial evidence for conservation of the pink-eyed dilution and uw genes in different classes of mammals including ungulates, lagomorphs, canines, marsupials, and primates (see below). For pink-eyed dilution, mice, deermice, rats, rabbits, and cats all have an equivalent locus, deﬁned by a tyrosinase-positive, OCA2-like phenotype, which is part of a conserved linkage group in these species (Heim et al., 1988; reviewed by Little, 1958). The porcine ortholog has been identiﬁed (Fernandez et al., 2002). In many mammalian species, the conserved linkage group includes the b-globin gene cluster. However, in humans, the b-globin locus (on human chromosome 11) is not linked to OCA2 (Heim et al., 1988), now known to be associated with mutations of the P gene on chromosome 15 (Durham-Pierre et al., 1994; Lee et al., 1994a, b; Rinchik et al., 1993). Like the mouse p gene, the chicken locus “pinkeye,” pk, causes a severe reduction in retinal pigmentation, produces gray feathers in strains in which wild-type birds produce black feathers, and shows histologic similarities to the melanocytes of p/p mice (Brumbaugh and Lee, 1975). The Medaka ﬁsh has a p gene ortholog (Fukamachi et al., 2004), and the p gene shares signiﬁcant homology with genes from Drosophila, yeast, and bacteria (Gardner et al., 1992; Rinchik et al., 1993).

The Mouse Matp Gene and Its Mutant Alleles Another gene likely to function in transport across the melanosome membrane is Matp (membrane-associated transport protein), previously known as AIM-1 (antigen in melanoma 1), a gene expressed exclusively in melanocytes and melanoma (Harada and Robbins, 1999; Harada et al., 2001; GenBank #AF172849). Note there are two other distinct genes called AIM-1 in the literature (Katayama et al., 1998; Teichmann et al., 1998). Among the mouse mutations that lead to severe hypopigmentation, without other phenotypes

235

CHAPTER 12

+

Amino acid transporter

+

Na H exchanger

Tyr (OCA1) Tyrp1 (OCA3)

H+

Na+ Tyrosine Melanin H+ H+ OR Anions Osmolyte Na+ Na+

H+/ATPase Fig. 12.2. Electron micrograph of choroid melanosomes of an adult mouse eye. Left: C57BL/6 (wild type) showing typically well-pigmented and round melanosomes. Right: mutant Matpuw-d/ Matpuw-d showing irregularly shaped melanosomes with uneven distribution of melanin. (Adult Matpuw-d/Matpuw-d mice have dark eyes.) Similar changes are seen in the RPE. Figures of eye melanocytes from these and other mutant Matp alleles are presented in Sweet et al. (1998).

(i.e. effects restricted to melanocytes), mutations in the Matp gene are distinguished by an unusual series of alleles (underwhite) with a spectrum of pigmentation phenotypes. Mutant Matp alleles are associated with small, crenated melanosomes with a range of pigment (Sweet et al., 1998). The original Matpuw allele arose spontaneously in the C57BL/6 strain and is recessive. The outer fur of Matpuw/Matpuw mice is a light beige with very white underfur, and the eyes are pink at birth, but darken with age (Dickie, 1964). Two other recessive alleles, Matpuw-i and Matpuw-d, have also been described (Sweet et al., 1998); homozygous Matpuw-i are slightly more pigmented than homozygous Matpuw; homozygous Matpuw-d are even more pigmented (brown/slate; Sweet et al., 1998). A fourth allele, MatpUw-dbr (dominant brown) is semi-dominant over the wild-type allele, and heterozygous MatpUw-dbr mice have a dark brown coat, while homozygous MatpUw-dbr mice are only slightly darker than uw/uw (Sweet et al., 1998). The phenotype of the eyes of uw allelic series was reported by Sweet et al. (1998), who noted normal numbers of melanocytes or melanosomes in the choroid or retinal pigmented epithelia in mice homozygous for mutant uw alleles. However, the melanosomes of both layers are generally smaller, irregularly shaped (raisin-like or crenated), and contain less pigment than those in wild-type eyes (Fig. 12.2). The action of the Matp gene is autonomous to melanocytes, as shown by transplantation assays (Lehman et al., 2000); its expression is an early marker of pigment cell precursors (Baxter and Pavan, 2002). Because Matp mutations lead to alterations in melanosome shape and pigmentation, it is likely that Matp is associated with the melanosome. The most severe mutant mouse allele, Matpuw/Matpuw is associated with a 7-bp deletion in the coding sequence of exon 3 (Du and Fisher, 2002) leading to a frameshift and resulting in an unstable transcript. Matp transcripts are expressed from the two other alleles that have intermediate pigmentation, but each mutant allele has a unique point mutation that leads to a nonconservative amino acid substitution (Matpuw-d: serine to proline at amino acid 435; MatpUw-dbr: aspartic acid to asparagine at amino acid 153), consistent with their partial function (Du and Fisher, 2002, Newton et al., 2001). 236

OR

P transporter (OCA2)

H+

Matp/uw transporter (OCA4)

Fig. 12.3. A simpliﬁed melanosome with key proteins. The melanosome organelle is the site of melanin biosynthesis and storage. The vacuolar H+/ATPase brings in protons (Dell’Angelica et al., 2000; Orlow, 1995; Smith et al., 2004). Two possible functions of the P transporter protein are presented [based on the function of homologous proteins: the P protein may facilitate anion transport needed to maintain electrogenic neutrality or it may function as an Na+/H+ antiporter (exchanger) (Puri et al., 2000)]. One or both Na+/H+ exchangers (NHE3 and/or NHE7) may also modulate the cations present in the melanosome (Smith et al., 2004). Consistent with its high homology to proton- and sodiumsugar symporters, the MATP protein may bring in a sugar (osmolyte) coupled with a proton or sodium (Newton et al., 2001). The model is consistent with changes in tyrosinase activity in melanosomes seen after treatment with agents that disrupt the pH gradient (Ancans et al., 2001; Halaban et al., 2002; Watabe et al., 2004), and differential timing of expression in melanogenesis allows for a dynamic change in the luminal pH. Thus, the P and Matp proteins play a major role in the normal regulation of melanosome biogenesis and melanin content. NB. The melanosome membrane location for Matp is based solely on the phenotype of mutant uw melanocytes. We also note that P protein has been reported in the ER and that both the P and the Matp proteins may primarily function in the vesicles that give rise to melanosomes, if not in the mature organelle.

The Matp Protein and Its Function Matp is transcriptionally modulated by MITF, the melanocytespeciﬁc transcription factor essential for pigmentation and a clinical diagnostic marker in human melanoma (Du and Fisher, 2002). Matp shares homology with known transporters, especially with plant H+/sucrose symporters (Newton et al., 2001). Indeed, the ﬁve amino acids that are conserved among all known plant H+/sucrose symporters are exactly conserved in mouse and human Matp. Plant H+/sucrose symporters couple the transport of a sucrose molecule with a proton along a proton gradient and often mediate changes in osmolarity (Stadler et al., 1999). Although there are no known mammalian sucrose transporters, it is possible that Matp transports a different sugar (or other related molecule) coupled with proton movement. Along with a reduction in pigment content,

MOLECULAR REGULATION OF MELANIN FORMATION: MELANOSOME TRANSPORTER PROTEINS

melanosomes from Matp mutant mice are small and irregularly shaped (raisin-like, crenated structures; Sweet et al., 1998; Fig. 12.2), consistent with a disruption in osmotic regulation. Moreover, drug- and sugar-mediated disruptions of the osmotic balance of endosomes and lysosomes have effects on protein turnover and luminal pH (Isaac et al., 2000; Schreiber and Haussinger, 1995; Schreiber et al. 1996), and the transport of amino acids across membranes is affected by osmotic imbalance (Gomez-Angelats et al., 1997) and pH (Potterf et al., 1996). Therefore, the Matp transporter is potentially an osmotic and/or pH regulator.

OCA4 and the Human Matp Gene Based on the mouse phenotype, MATP was evaluated as a candidate gene for a form of OCA with a phenotype similar to OCA2 (P gene albinism). The MATP gene was sequenced in several OCA individuals who had minimal pigment, a few of whom had previously been screened for mutations on the P gene. One Turkish OCA individual (with distantly related parents) was found to be homozygous for a splice site mutation of the MATP gene (G to A in the splice acceptor sequence of exon 2) and did not have a P gene mutation. This patient deﬁned a new form of OCA, OCA4 (Newton et al., 2001). Subsequently, MATP mutations leading to OCA4 have been described in additional Turkish individuals (264delC; Ikinciogullari et al., 2004) and several German individuals (L361P, P58A, Y401X, F221del, T329fsX68, Y317C, A477T, W202C, F525fsX15; IVS3+46C4T, IVS3+59+61dup; Rundshagen et al., 2004). In addition, OCA4 is one of the most common forms of albinism in the Japanese population, where 24% of OCA is OCA4, associated with seven mutant alleles [four missense mutations (P58S, D157N, G188V, and V507L) and three frameshift mutations (S90CGGCCAÆGC, V144insAAGT, and V469delG); Inagaki et al., 2004].

Evolution of the Matp Gene The cream color in horses has been associated with the same mutation as the MatpUw-dbr mouse (D153N; Mariat et al., 2003). Mutations of the Medaka ortholog (the B gene) have been identiﬁed in the Medaka ﬁsh (Fukamachi et al., 2001). The Matp gene shares signiﬁcant homology with genes from Drosophila and, as mentioned earlier, genes as distant as plant proton/sucrose symporters.

Other Proteins Involved in Transport Across the Melanosome Membrane In addition to the p protein and Matp, other proteins may also play a role in ion transport across melanosome membranes. Recently, two Na+/H+ exchangers, NHE3 and NHE7, have been found associated with melanosomes, and the expression of these Na+/H+ exchangers may vary between Caucasian and black melanocytes (Smith et al., 2004), suggesting that other proteins in addition to p and Matp may regulate ion ﬂux and/or pH, playing a role in the regulation of melanin synthesis in melanosomes. These researchers also conﬁrmed that the melanosome has an ATP-driven proton pump.

Combining what we know about the phenotypes of mutant p and uw melanocytes, the functions of proteins sharing structural similarity with the p and Matp proteins, and the function of other transport proteins known to reside on the melanosome membrane, a model of melanosome transport is presented (Fig. 12.3).

Normal Human Pigmentation Variation and the P and MATP Genes Humans vary considerably in the range of normal pigmentation of the hair, eyes, and skin. It is reasonable to believe that at least part of this variation results from sequence variation in key pigmentation genes, such as the P and MATP genes that underlie forms of albinism. This notion is supported by the range in pigmentation phenotypes seen among mutations of the mouse and human orthologs. Mouse phenotypes vary among different p alleles (Lyon et al., 1992; Russell et al., 1995), and a wide range in OCA2 phenotypes has been noted for various P alleles (Oetting and King, 1999; see Chapter 31). There is also a wide range of phenotypes among mice with various mutant Matpuw alleles (Sweet et al., 1998), and among the limited number OCA4 patients examined to date (Inagaki et al., 2004; Newton et al., 2001; Rundshagen et al., 2004). A gene associated with brown eyes (total brown iris pigment) and a gene associated with brown hair map to 15q, with the P gene as the prime candidate (Eiberg and Mohr, 1996). Population studies looking at the association of speciﬁc polymorphisms of the P and MATP genes demonstrate that (together with polymorphisms of the MC1R gene) they play a major role in determining the normal range of pigmentation of the hair (Eiberg and Mohr, 1996, Sturm et al., 2001), skin (Akey et al., 2001; Duffy et al. 2004; Nakayama et al., 2002; Shriver et al., 2003), and eyes (Frudakis et al., 2003; Rebbeck et al., 2002; Sturm and Frudakis, 2004; Zhu et al., 2004). Indeed, a polymorphism of MATP (L374F) may be a useful marker of population origin (Yuasa et al., 2004).

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The Pigmentary System: Physiology and Pathophysiology, Second Edition Edited by James J. Nordlund, Raymond E. Boissy, Vincent J. Hearing, Richard A. King, William S. Oetting, Jean-Paul Ortonne Copyright © 2006 Blackwell Publishing Ltd

13

Transcriptional Regulation of Melanocyte Function Kazuhisa Takeda and Shigeki Shibahara

Summary 1 Microphthalmia-associated transcription factor (Mitf) encoded at the mouse microphthalmia locus or its human homolog MITF is a cell type-speciﬁc regulator that is required for development and/or survival of pigment cells. Heterozygous mutations in the MITF gene cause auditory–pigmentary syndromes, known as Waardenburg syndrome type 2. 2 MITF consists of at least seven isoforms, such as MITF-A, MITF-B, MITF-C, MITF-H, and MITF-M, differing at their N-termini and in their expression proﬁles. MITF-M is a founder member of multiple MITF isoforms and represents a melanocyte-speciﬁc isoform. Each N-terminal region is encoded by a separate exon 1 of the MITF gene. 3 The melanocyte-speciﬁc (M) promoter of the MITF gene is upregulated by several transcription factors, PAX3, SOX10, LEF-1, and CREB. The activity and degradation of MITF-M are also regulated by extracellular signals via protein phosphorylation, such as c-Kit signaling. Together, multiple signals appear to converge on the M promoter as well as on the MITF-M protein. 4 The tyrosinase family consists of tyrosinase, tyrosinaserelated protein 1 (TRP-1), and TRP-2/DOPAchrome tautomerase. We have summarized the short history of identiﬁcation and characterization of these proteins, which are speciﬁcally or preferentially expressed in pigment cells. The cis-regulatory elements of the tyrosinase family genes have been identiﬁed in the 5¢ ﬂanking region of each gene. Many of these regulatory elements contain the CATGTG motif, a consensus sequence recognized by a large family of transcription factors with a basic helix–loop–helix structure, such as Mitf. 5 Retinal pigment epithelium (RPE) is derived from the optic cup of embryonic brain and shares the capacity of melanin production with melanocytes. RPE forms a single layer of cells interposed between the neural retina and the vascular-rich choroids, thereby constituting the blood–retinal barrier. RPE is responsible for correct development of the eye and for survival of photoreceptors in adult retinas.

Historical Background Pigmentation is one of the most demonstrable phenotypes in the animal kingdom, and melanin is a principal pigment found 242

in all living organisms, including fungi and plants. In the mouse, at least 90 different loci have been known to affect coat color. Thus, mice coat color mutations provide an excellent model for studying gene action and its regulation. Melanin production is speciﬁcally seen in the differentiated melanocytes that originate from the neural crest and in the RPE derived from the optic cup of the brain. Like other cell lineages of neural crest origin, melanoblasts, a precursor to melanocytes, migrate throughout the dermis during development. The RPE, however, does not need to migrate such a long distance during development. The RPE is a monolayer epithelium interposed between the vascular-rich choroid and the neural retina, and such an anatomical organization of RPE is entirely different from that of melanocytes. In spite of these differences, both cell types share the unique capacity of melanin production. Tyrosinase (EC 1.14.18.1) is a rate-limiting enzyme in melanin biosynthesis and has been a symbolic enzyme in pigment cell research. Thus, using various approaches, many investigators have attempted to purify and characterize tyrosinase. The ﬁrst pigment cell-speciﬁc cDNA was isolated by differential hybridization from B16 mouse melanoma cells and was initially suggested to encode tyrosinase (Shibahara et al., 1986). This report deﬁnitely facilitated the research related to pigment cells, although the cDNA has been shown not to encode tyrosinase but what is now known as tyrosinaserelated protein 1 (TRP-1) (Jackson, 1988). Since then, many genes related to pigmentation, including tyrosinase, have been cloned. In particular, the second enzyme structurally related to tyrosinase was discovered and termed tyrosinase-related protein 2 (TRP-2) (Jackson et al., 1992; Tsukamoto et al., 1992). There is about 40% amino acid identity among tyrosinase, TRP-1, and TRP-2, all of which constitute the tyrosinase family and are speciﬁcally expressed in melaninproducing cells. TRP-2 possesses the activity of DOPAchrome tautomerase (EC 5.3.2.3), catalyzing the conversion of DOPAchrome to 5,6-dihydroxyindole-2-carboxylic acid (DHICA) (Kroumpouzos et al., 1994; Yokoyama et al., 1994a). TRP-2 is now known as DOPAchrome tautomerase (DCT). TRP-1 was shown to catalyze the conversion of DHICA to indole–quinone–carboxylic acid (JiménezCervantes et al., 1994; Kobayashi et al., 1994). Thus, both TRP-1 and DCT/TRP-2 are the enzymes that determine the quality of melanin. The tyrosinase gene family has been characterized as a model to study the regulation of pigment cellspeciﬁc and developmental stage-speciﬁc gene transcription.

TRANSCRIPTIONAL REGULATION OF MELANOCYTE FUNCTION ∆R217 A→C in splice acceptor (Tietz) R259Stop G→A in splice donor R214Stop N278D S250P S298P

AE

MITF

mirw

miws

bHLH

misp Miwh

LZ

mi Mior

mivit

Internal Cytosine I212N ∆R216 R216K D222N Deregulation and novel exon 1 deletion insertion miew Mib ∆exons 2–4 in splice acceptor 2 Internal G244E deletion ∆exon 6

mice R263Stop

Fig. 13.1. Schematic illustration of Mitf/MITF and the characterized mutations. A melanocyte-speciﬁc isoform, Mitf-M/MITF-M, is shown. The nine exons are indicated by alternate open and closed boxes over the protein structure. The mutations found in individuals affected by WS2 syndrome (Nobukuni et al., 1996; Tassabehji et al., 1994, 1995) are shown above the functional domains. The delR217 is responsible for Tietz syndrome (Amiel et al., 1998). Asterisks and triangles indicate the splicing mutations and the nonsense mutations respectively. The mutations at the 10 Mitf alleles are indicated below (Hodgkinson et al., 1993; Steingrímsson et al., 1994, 1996), and the mutant alleles shown within boxes are unable to bind to DNA in vitro (Hemesath et al., 1994). Region AE indicates the activating exon (exon 4).

Here, we summarize recent advances concerning mechanisms for pigment cell-speciﬁc transcription of the tyrosinase family genes. Elucidation of the mechanisms regulating transcription of the tyrosinase family genes is an essential requirement for understanding the molecular events underlying differentiation of pigment cells. A comprehensive text (Silvers, 1979) and good reviews on mouse coat color mutations have been published (Jackson, 1994; Jackson et al., 1994; Urabe et al., 1993).

Genes Regulating the Development of Pigment Cells The genes described in this section are required for development and migration of melanoblasts and melanocytes, and the mutations of each gene cause white spotting or entirely white coat color, characterized by the lack of melanocytes. Most of these mutations do not affect the development of RPE, except for the microphthalmia-associated transcription factor (Mitf); Mitf is required for normal development of neural crestderived melanocytes and RPE.

Microphthalmia-Associated Transcription Factor Overview Mice with mutations at the microphthalmia (mi) locus, now termed Microphthalmia-associated transcription factor (Mitf), have some or all of the following defects: loss of pigmentation, reduced eye size, failure of secondary bone resorption, reduced numbers of mast cells, and early onset of deafness (Silvers, 1979). The most severe manifestations of Mitf alleles, the original Mitf mi and Mitf or (microphthalmia Oak Ridge), are inherited semi-dominantly and include microphthalmia, white coat, deafness (due to lack of inner ear melanocytes),

and osteopetrosis. Less severe alleles tend to be recessive. These phenotypic variabilities have been shown to represent allelic differences. All Mitf alleles are characterized by defects in neural crest-derived melanocytes, leading to coat color dilution, white spotting or complete loss of pigmentation, and deafness. It is clear that the Mitf protein encoded at the Mitf locus plays an essential role in development and/or survival of several cell lineages, including RPE, mast cells, and osteoclasts. Mitf protein has been shown to play a role in promoting the transition of precursor cells to melanoblasts and melanoblast survival (Opdecamp et al., 1997). Mitf as a Member of a Subfamily of Transcription Factors with a bHLH-LZ Independent transgenic insertions disrupted the Mitf locus on mouse chromosome 6 (Krakowsky et al., 1993; Tachibana et al., 1992), which allowed two groups to clone a novel factor encoded at the Mitf locus (Hodgkinson et al., 1993; Hughes et al., 1993). Mitf consists of 419 amino acid residues and is predicted to contain a basic helix–loop–helix and a leucine-zipper (bHLH-LZ) structure (Fig. 13.1). This type of Mitf is now known as Mitf-M representing a melanocyte-speciﬁc isoform (Amae et al., 1998). The bHLH-LZ structure is required for DNA binding and dimerization (reviewed by Kadesch, 1993). The predicted Mitf protein shows the highest degree of amino acid similarity to TFE3 and TFEB, both of which are ubiquitously expressed and bind the palindromic sequence (CACGTG), either as homodimers or as TFE3–TFEB heterodimers (Fisher et al., 1991). TFE3 was originally identiﬁed as a human protein that binds the E3 box of the m chain immunoglobulin enhancer (Beckmann et al., 1990), and TFEB as the human protein that binds to the adenovirus major late transcription factor (Carr and Sharp, 1990). Hemesath et al. (1994) have shown that Mitf binds in vitro to DNA 243

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containing the CACGTG motif derived from adenovirus major late promoter as a homo- or heterodimer with TFEB, TFE3, or TFEC. TFEC was originally identiﬁed as a bHLH-LZ protein, forms heterodimers with TFE3, and inhibits TFE3dependent transcription activation (Zhao et al., 1993). TFEC also shows a high degree of amino acid sequence similarity to MITF (Yasumoto and Shibahara, 1997). In contrast, no heterodimers were observed upon mixing Mitf with Max, Myc, or USF. Mitf also bound the CATGTG motif, a core element of the M-box, and was shown to trans-activate the Mbox-driven reporter gene in ﬁbroblasts (Hemesath et al., 1994). Nakayama et al. (1998) reported that expression of TFEB or TFE3 is undetectable in the neural crest at stages when the Mitf mutant phenotype becomes manifest. Furthermore, the phenotypes of Mitf mutant mice in melanocytes and RPE were not affected in double and triple mutant mice lacking Mitf, TFE3, and/or TFEC (Steingrímsson et al., 2002). These results suggest that Mitf may function as a homodimer during the development of melanocytes and RPE. The human homolog of the mouse Mitf gene, MITF, was cloned and mapped to chromosome 3p12.3–p14.1 (Tachibana et al., 1994). MITF-M, a melanocyte-speciﬁc isoform, is predicted to consist of 419 amino acid residues and share 94.4% identity with mouse Mitf-M. The predominant species of MITF mRNA is of about 5.5 kb in human melanoma cells (Tachibana et al., 1994). Ectopic expression of MITF-M converts ﬁbroblasts to cells with melanocyte characteristics (Tachibana et al., 1996). Multiple Isoforms of Mitf with Distinct Amino-Termini The MITF gene is expressed as multiple isoforms in humans and mice, termed MITF-M, MITF-A, MITF-B, MITF-C, MITF-D, MITF-E, MITF-H, and MITF-mc (Amae et al., 1998; Fuse et al., 1999; Oboki et al., 2002; Steingrímsson et al., 1994; Takeda et al., 2002; Takemoto et al., 2002; Yasumoto et al., 1994; for a review, see Shibahara et al., 2001). These isoforms share common bHLH-LZ and transcription activation domains, but differ in the amino-terminal regions (Fig. 13.2). MITF-M is a founder member of multiple MITF isoforms, and its amino-terminus, domain M, consists of 11 amino acid residues and is encoded by the exon 1M. All other isoforms with the extended amino-termini share the entire carboxyl portion with MITF-M. The unique aminoterminus of MITF-A, MITF-B, MITF-C, MITF-MC, or MITFH shares the common region of 83 amino acid residues (domain B1b), which is signiﬁcantly similar to the equivalent portion of TFEB (Amae et al., 1998) and TFE3 (Rehli et al., 1999; Yasumoto et al., 1998). The initiating methionine of MITF-D or MITF-E is located in the B1b domain, as each exon 1 is a noncoding exon. The MITF gene consists of many promoters, their consecutive ﬁrst exons, such as exon 1A, 1H, 1D, 1B, and 1M (Fig. 13.2), and eight downstream exons that are common to all isoforms (Hallson et al., 2000; Udono et al., 2000). The unique amino-termini of the MITF isoforms are encoded by separate ﬁrst exons, except for exon 1D, which codes for the 244

Deletion in Mitf mi-rw 1A

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Fig. 13.2. Structural organization of the MITF gene encoding multiple isoforms. The protein-coding regions are indicated by closed boxes, and the 5¢- and 3¢-untranslated regions are indicated by open boxes. Each arrow indicates the transcription start site of separate exon 1. MDE represents the MITF-M distal enhancer. The Mitf mi-rw gene lacks the DNA segment containing exon 1H and MDE (Hallsson et al., 2000; Watanabe et al., 2002). The Mitf mi-bw gene contains the L1 element insertion in intron 3 (Yajima et al., 1999). The bottom panel shows the representative MITF isoforms, MITF-M, MITF-A, MITF-H, and MITF-D. Region A indicates the transcription activation domain. Regions Thr and Ser represent the threonine-rich domain and the serine-rich domain respectively.

5¢ untranslated region of the mRNA. Among the ﬁrst exons, exon 1B is unique because it is also used as a second exon (B1b exon), when the primary transcripts, initiated from exon 1A, exon 1H, or exon 1D, are subjected to splicing (Udono et al., 2000). Exon 1B therefore encodes the 5¢ untranslated region of MITF-B mRNA, the amino-terminal domain of MITF-B (domain B1a), and domain B1b (Fig. 13.2). Dysfunction of Mitf impairs the differentiation and/or survival of neural crest-derived melanocytes in skin and ear, and optic cup-derived RPE in the eye (Hodgkinson et al., 1993). Among the MITF isoforms, MITF-M is exclusively expressed in melanocytes and pigmented melanoma cells, but is not detectable in any other cell types, including human RPE cell lines (Amae et al., 1998; Fuse et al., 1999). In contrast to MITF-M, other MITF isoforms are widely expressed in many cell types including human RPE cell lines (Amae et al., 1998; Fuse et al., 1999; Takeda et al., 2002; Udono et al., 2000). The Mitf gene is also expressed in heart (Steingrímsson et al., 1994) and male germ cells (Saito et al., 2003a), but no abnormalities in these tissues have been described in Mitf mutants. Informative Mitf Alleles The Mitf gene is altered in two independent Mitf alleles, Mitf mi and Mitf ws (Hodgkinson et al., 1993). Both alleles are semidominant, but the original Mitf m allele shows a more severe phenotype. The Mitf mi allele contains a three-base deletion, resulting in loss of an arginine residue from a series of four consecutive conserved arginines (residues 214–217), located at the carboxyl end of the DNA-binding basic region, whereas the Mitf ws mutation results from an intragenic deletion

TRANSCRIPTIONAL REGULATION OF MELANOCYTE FUNCTION

(Fig. 13.1). Subsequently, it was shown that a protein encoded at the Mitf mi allele is unable to bind in vitro to the CACGTG motif (Hemesath et al., 1994) and to the CATGTG motif (Hemesath et al., 1994; Morii et al., 1994). The deletion of the Mtﬁ gene was also found in genomic DNA of two Mitf alleles, Mitf ws and Mitf rw (Hughes et al., 1993). Molecular defects associated with eight murine Mitf mutations were characterized (Steingrímsson et al., 1994), and the DNA-binding abilities of mutant proteins were examined in vitro (Hemesath et al., 1994), clearly indicating that the phenotypic variation associated with various Mitf alleles can be explained in large part by the underlying nature of the mutation. For example, a mutation associated with a semi-dominant allele, Mitf or, is an Arg216Lys substitution in the basic region (Steingrímsson et al., 1994), where a deletion of one arginine residue of the four consecutive arginine residues (214–217) is present in the Mitf mi mutation (Hodgkinson et al., 1993) (Fig. 13.1). Indeed, both Mitf mi and Mitf or produce nearly identical phenotypes, including osteopetrosis, which represents the defect of bone resorption and is caused by the osteoclast dysfunction. It was then conﬁrmed that the mutation associated with Mitf mi or Mitf or alleles completely abolishes DNA-binding ability as homodimers or heterodimers with TFE3 in vitro (Hemesath et al., 1994). In addition, the presence of either Mitf mi or Mitf or protein ablated TFE3 homodimeric DNA-binding activity. Thus, the semi-dominant Mitf mutations may represent dominant-negative mutations, as each mutation is predicted to inhibit Mitf function. It has also been reported that the nuclear translocation potential of Mitf mi and Mitf ew proteins is impaired and thus, unlike the nuclear-located Mitf protein, both mutant proteins are predominantly located in the cytoplasm (Takebayashi et al., 1996). Essential requirement of Mitf-M for melanocyte development was veriﬁed by the molecular lesion of black-eyed white Mitf mi-bw mice (Yajima et al., 1999), which are characterized by the complete white coat color, deafness, and normally pigmented RPE without any ocular abnormalities. In Mitf mi-bw mice, the insertion of an L1 retrotransposable element in the intron 3 between exon 3 and exon 4 leads to complete repression of Mitf-M mRNA expression and to reduction in Mitf-A and Mitf-H mRNAs expression (Yajima et al., 1999). In this context, the M promoter represents the most downstream promoter of the MITF/Mitf gene (Fuse et al., 1996; Hallsson et al., 2000; Tassabehji et al., 1994; Udono et al., 2000), and may be most susceptible to the transcriptional repression caused by the insertion of the L1 element (Fig. 13.2). These results indicate that MITF-M/Mitf-M is a key regulator of melanocyte development but is dispensable for RPE development. The homozygous red-eyed white Mitf mi-rw mice exhibit small red eyes and white coat with some pigmented spots around the head and/or tail (Steingrímsson et al., 1994), and its molecular defect is a deletion of the genomic DNA segment containing exon 1H and exon 1B (Hallsson et al., 2000) (Fig. 13.2). It is therefore conceivable that the phenotype of Mitf mi-rw mice represents the loss of function of all Mitf isoforms with extended amino-termini. Moreover, these mutant

mice are deﬁcient in melanocytes, probably due to the loss of Mitf-M expression, except for melanocytes located in the head and tail regions, suggesting that the deleted genomic DNA segment may contain the enhancer for the M promoter. In fact, an upstream enhancer has been identiﬁed, termed MDE (Fig. 13.2) (Watanabe et al., 2002). Mutations in the MITF Gene, Causing Auditory–Pigmentary Syndrome Waardenburg syndrome type 2 (WS2) is an autosomal dominant disorder of sensorineural hearing loss and pigmentary disturbances, and its responsible gene has been mapped to chromosome 3p12.3–p14.1, close to the position of the MITF gene (Hughes et al., 1994). Subsequently, mutations affecting splice sites in the MITF gene were identiﬁed in two families with WS2 (Tassabehji et al., 1994). MITF gene dosage may be more critical in the developing human than in the mouse. A mutation responsible for Tietz syndrome (albinism– deafness syndrome) has been identiﬁed as an in-frame deletion of the MITF gene, removing one of four consecutive Arg residues (delR217) in the basic region of MITF-M (Amiel et al., 1998) (Fig. 13.1). Tietz syndrome is characterized by profound deafness, generalized albinism with blue eyes, and hypoplasia of the eyebrows. Unlike WS2, Tietz syndrome shows dominant inheritance with complete penetrance and does not involve the eyes. Thus, the delR217 mutation causes more severe dysfunction of melanocytes than do other MITF mutations. Notably, the delR217 mutation found in Tietz syndrome is equivalent to the mouse semi-dominant Mitf mi mutation (Hodgkinson et al., 1993). The functional consequence of the delR217 mutation was shown to be a loss of DNA-binding activity and to function as a dominant-negative form of MITF-M (Hemesath et al., 1994). In contrast to the haploinsufﬁciency of MITF in WS2 (Saito et al., 2002), a dominant-negative function of the delR217 MITF-M may be responsible for Tietz syndrome. Regulation of the MITF-M Promoter by Multiple Transcription Factors MITF-M expression is upregulated by Wnt signaling (Dorsky et al., 2000; Takeda et al., 2000a). Wnt proteins, secreted cysteine-rich glycoproteins, have been established as developmentally important signaling molecules (reviewed by Cadigan and Nusse, 1997). Wnt signaling induces intracellular accumulation of b-catenin, which functions as a coactivator for LEF-1/TCF transcription factors (reviewed by Cadigan and Nusse, 1997; Clevers and van de Wetering, 1997; Eastman and Grosschedl, 1999). A double knockout mouse of Wnt-1 and Wnt-3a genes exhibits deﬁciency of neural crest derivatives, including melanocytes (Ikeya et al., 1997). Wnt signaling promotes pigment cell formation from neural crest in zebraﬁsh (Dorsky et al., 1998). Moreover, the onset of Wnt3a expression is detected at 7.5 embryonic days (Takada et. al., 1994), which precedes the onset of Mitf expression in neural crest cells (9.5–10.5 embryonic days) (Nakayama et al., 1998). Thus, MITF-M is a good candidate for the target genes of Wnt signaling. Indeed, the MITF-M promoter is bound by 245

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LEF-1 and is activated by Wnt-3a (Takeda et al., 2000a). Wnt3a protein induces endogenous Mitf-M mRNA in cultured melanocytes (Takeda et al., 2000a). Interestingly, the MITFM by itself interacts with LEF-1, which efﬁciently activates its own transcription (Saito et al., 2002; for a review, see Saito et al., 2003b). Recruitment of MITF-M on the M promoter is an essential step for the self-activation of MITF-M expression and depends on the binding of LEF-1 to the three adjacent binding sites (Saito et al., 2002). It is therefore likely that transcription from the M promoter is relatively sensitive to the concentrations of LEF-1 and MITF-M. Importantly, MITF-M is able to transactivate the M promoter by interacting with LEF-1 but without binding to the M promoter, because the M promoter does not contain the CATGTG motif and is not bound by MITF-M in vitro. In addition, the mutant MITF-M protein lacking the DNA-binding activity also enhances the LEF-1mediated transactivation of the M promoter (Saito et al., 2002). We used the Gly244Glu substitution found in the semidominant Mitf brownish (Mitf b) mouse (Steingrímsson et al., 1996). Likewise, Mitf-M has been shown to interact with c-jun to transactivate the mouse mast cell protease 7 gene promoter that lacks a typical MITF-binding element (Ogihara et al., 2001). It is therefore conceivable that MITF-M functions as a nonDNA-binding cofactor for LEF-1 on the M promoter. This notion is of physiological signiﬁcance in understanding the phenotypic consequences of various MITF and Mitf mutations that alter the DNA-binding activity. MITF-M expression is also upregulated by a-melanocytestimulating hormone (a-MSH), which stimulates cAMP signaling and activates the transcription factor CREB by phosphorylation. CREB binds the cAMP-responsive element in the MITF-M promoter, resulting in transcriptional activation of the MITF-M gene (Price et al., 1998). The cAMP signalingmediated activation of the MITF-M gene is melanocyte speciﬁc (Price et al., 1998). This activation requires SOX10, a transcription factor containing a high-mobility-group domain (Huber et al., 2003). The Function of MITF-M Protein is Regulated by Phosphorylation The Steel/c-kit signaling triggers phosphorylation of MITF-M via the MAP kinase ERK2 at Ser-73, and upregulates the function of MITF-M (Hemesath et al., 1998). The function of MITF-M protein is regulated by many signals, which induce phosphorylation of MITF-M (see Fig. 13.4). Consequently, MITF-M could interact with CBP/p300 transcriptional coactivator efﬁciently (Price et al., 1998; Sato et al., 1997). Moreover, c-kit signaling results in phosphorylation of MITF-M at Ser-409 via p90 Rsk, a member of the serine/threonine kinases, and causes short-lived activation and destruction of MITF-M (Wu et al., 2000). In this context, the ubiquitin-conjugating enzyme hUBC9 was identiﬁed as a potential interacting partner for MITF-M (Xu et al., 2000). It was shown that phosphorylation of MITF-M at Ser-73 is a prerequisite for the hUBC9-mediated degradation of MITF-M. 246

Glycogen synthase kinase 3b (GSK3b) phosphorylates MITF-M at Ser-298, enhancing the DNA-binding ability of MITF-M (Takeda et al., 2000b). In fact, a Ser298Pro substitution was found in the individuals affected by WS2 (Tassabehji et al., 1995). It has been reported that GSK3b is activated by cAMP, which stimulates melanogenesis (Khaled et al., 2002). Mitf Regulates the Genes Involved in Melanin Formation and Melanocyte Survival Mitf protein transactivated the mouse tyrosinase (Ganss et al., 1994a) and TRP-1 promoters probably by binding to the Mbox (Yavuzer et al., 1995). It was also shown that Mitf protein activates the human tyrosinase promoter mainly through the initiator E-box (Bentley et al., 1994). Likewise, MITF noticeably increased the expression of a reporter gene under the control of the human tyrosinase promoter, mainly through the CATGTG motif of the tyrosinase distal element, TDE (positions –1861 to –1842) (Yasumoto et al., 1994). Subsequently, MITF was shown to bind in vitro to TDE and was also shown to transactivate the mouse tyrosinase and TRP-1 promoters, probably by binding to the M-box (Yasumoto et al., 1995). The M-box was originally identiﬁed as a positive regulatory element in the mouse TRP-1 promoter (Lowings et al., 1992) and is well conserved in the tyrosinase family genes. In contrast, although DOPAchrome tautomerase (DCT) promoter contains the M-box, its promoter is not activated by MITFM alone (Yasumoto et al., 1997). Subsequently, it has been reported that the DCT promoter is activated by cooperation of MITF with LEF-1 (Yasumoto et al., 2002). These results indicate that MITF is a factor required for pigment cellspeciﬁc transcription of the tyrosinase family genes. It is noteworthy that MITF may form a dimer with a cell-speciﬁc or ubiquitous DNA-binding protein, which in turn functions to direct the pigment cell-speciﬁc transcription of the tyrosinase family genes. Other pigment cell-speciﬁc target genes of MITF are Tbx2, a member of the T-box family (Carreira et al., 2000), melanosomal proteins Pmel17 (silver/gp100) and MART1/ MELANA (Du et al., 2003), and MELASTATIN (MLSN1/ TRPM1), an ion channel of the transient receptor potential (TRP) superfamily (Miller et al., 2004). MITF is also responsible for melanocyte survival by regulating the antiapoptotic gene Bcl2 (McGill et al., 2002), disruption of which causes depigmentation in second hair follicle change (Yamamura et al., 1996). Moreover, it has been shown that MITF interacts with the retinoblastoma product in vitro (Yavuzer et al., 1995). Thus, MITF may regulate the cell cycle, but it is unknown whether MITF transactivates the cell cycle-related genes.

Pax-3 or Splotch Homozygous splotch (Sp) mice usually die at 13 days of gestation with multiple abnormalities, including neural tube defects and dysgenesis of neural crest-derived cells (Auerbach, 1954; Silvers, 1979), whereas heterozygous splotch mice display white spotting on the belly and occasionally on the back, feet,

TRANSCRIPTIONAL REGULATION OF MELANOCYTE FUNCTION

and tail. It has been shown that the splotch mutation disrupts the development of functioning melanocytes but does not affect RPE (Auerbach, 1954; Pavan and Tilghman, 1994). The splotch locus encodes the Pax-3 protein, a member of a family of putative transcription factors related to the paired box family of Drosophila segmentation genes (Epstein et al., 1991). In the mouse, the Pax gene family encodes a group of eight related proteins, which share a domain homologous to that encoded by the Drosophila paired box (reviewed by Chalepakis et al., 1993). One mutant splotch allele, Sp2H, contains a deletion of 32 bp in the exon sequence coding for the paired homeodomain of Pax-3 protein (Epstein et al., 1991). Human PAX3 gene encodes a paired domain transcription factor located at chromosome 2q35, and its mutations are associated with Waardenburg syndrome type 1 (WS1) and type 3 (WS3) (Baldwin et al., 1992; Tassabehji et al., 1992, 1993, 1995). WS1 and WS3 are dominantly inherited syndromes of hearing loss, pigmentary disturbances, and heterochromia iridis. Unlike WS2, WS1 and WS3 are characterized by dystopia canthorum and limb abnormality, respectively. Watanabe et al. (1998) has shown that the MITF-M promoter is a direct target gene for PAX3, which accounts for the fact that mutations of either transcription factor PAX3 or MITF could cause Waardenburg syndrome. TRP-1 gene is also activated by PAX3 (Galibert et al., 1999).

SOX10 SOX10 is a transcription factor containing a high-mobilitygroup box as a DNA-binding motif, and is responsible for Waardenburg–Hirschsprung syndrome, also known as WS4 (Pingault et al., 1998), which is characterized by aganglionic colon, sensorineural deafness, and pigmentation abnormalities. In the spontaneous mouse mutant Dominant megacolon (Dom), Sox10Dom is functionally inactive due to a frameshift mutation (Herbarth et al., 1998; Southard-Smith et al., 1998). The Dom mice suffer from several neural crest defects including the enteric nervous system and melanoblasts. At E11.5, mouse embryos homozygous for the Sox10Dom mutation entirely lack neural crest-derived cells expressing the lineage marker DOPAchrome tautomerase (DCT) or Mitf. Moreover, neural crest cell cultures derived from homozygous embryos do not give rise to pigmented cells (Potterf et al., 2001). Thus, Sox10 may regulate transcription of the DCT and MITF genes. In the heterozygous Dom mice, c-kit-expressing melanoblasts are observed at E11.5, but lack Dct expression, suggesting that Sox10 may serve as a transcriptional activator of Dct. Indeed, Sox10 and Dct colocalized in early melanoblasts, and Sox10 is capable of transactivating the Dct promoter in a transient expression assay (Potterf et al., 2001). Moreover, Ludwig et al. (2004) have shown the synergistic activation of the Dct gene by Sox10 and Mitf. By in situ hybridization analysis, we have shown the coexpression of Sox10 and Mitf (most likely Mitf-M) mRNAs in migrating melanoblasts at E11.5 (Watanabe et al., 2000). By E13.5, Sox10 mRNA expression became undetectable in

migrating melanoblasts, whereas Mitf expression continues to be detectable. In postnatal day 8 and adult cochleas, Sox10 expression was detected only in the supporting cells of the organ of Corti (Watanabe et al., 2000). Taken together, these results support the notion that Sox10 is required for transcription from the M promoter of the MITF/Mitf gene during the early stage of melanoblast development. MITF-M gene is also activated by SOX10 via at least three SOX10 binding sites of the proximal promoter (Bondurand et al., 2000; Lee et al., 2000; Potterf et al., 2000; Verastegui et al., 2000). SOX10 synergistically activates the MITF-M promoter by cooperating with PAX3 (Bondurand et al., 2000; Potterf et al., 2000). SOX10 also binds the MITF-M distal enhancer (MDE) located at about the –15-kb region upstream of exon 1M (Watanabe et al., 2002). SOX10 was shown to function as an architectural transcription factor by bending the DNA (Peirano and Wegner, 2000). It is therefore conceivable that the DNA bending, induced by SOX10 bound to MDE, may facilitate the interaction with the transcription factors on the proximal MITF-M promoter region (Fig. 13.3). These data support a model in which deafness and hypopigmentation common to all types of WS are caused by a functional disruption of a key transcription factor, MITF-M.

CITED1 CITED1 was identiﬁed as a melanocyte-speciﬁc gene 1 (msg1), which is a nuclear protein expressed in pigmented melanocytes and melanoma cells, but not in amelanotic melanoma cells (Shioda et al., 1996). The normal tissue distribution of CITED1 in adult mice was conﬁned to melanocytes and testis. Murine Cited1 and human CITED1 genes encode a predicted protein of 27 kDa with 75% overall Wnt 5’ (MDE) –15 kb SOX10

Frizzled

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Fig. 13.3. Transcriptional regulation of the MITF-M promoter. The proximal MITF-M promoter is bound by PAX3, which is responsible for WS1 and WS3, and by SOX10, which is responsible for WS4. SOX10 also binds the MITF-M distal enhancer (MDE) to activate the MITF-M gene. Wnt signaling induces intracellular accumulation of b-catenin, which functions as a coactivator for LEF-1, leading to activation of the MITF-M gene. MITF-M also interacts with LEF-1 and activates its own transcription. a-Melanocyte-stimulating hormone (a-MSH) stimulates cAMP signaling and activates the transcription factor CREB by phosphorylation. CREB induces MITF-M mRNA expression via a cAMP-responsive element in the MITF-M promoter (Price et al., 1998).

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also detected in a mouse melanoblast cell line (Eisen et al., 1995; Thomson et al., 1993). Expression of BRN2, in melanoma cells in cotransfection assays, repressed the activity of the melanocyte-speciﬁc tyrosinase promoter (Eisen et al., 1995). Additionally, BRN2 ablated melanoma cells revert to a less mature cell type lacking many markers of differentiated melanocytes such as MITF (Thomson et al., 1995). These results suggest that BRN2 maintains the undifferentiated melanocytic phenotype.

Upstream Stimulatory Factor 1 (USF1)

Fig. 13.4. Post-translational regulation of MITF-M by multiple signals. Steel/c-kit signaling triggers phosphorylation of MITF-M via the MAP kinase ERK2 at Ser-73, and increases interaction of MITFM with CBP/p300 transcriptional coactivators. The c-kit signaling phosphorylates MITF-M at Ser-409 via a serine/threonine kinase p90 Rsk, and causes short-lived activation and destruction of MITF-M. For the degradation of MITF-M, the ubiquitinconjugating enzyme hUBC9 interacts with MITF-M. Endothelin 3, which is responsible for WS4, stimulates MAP kinase, and probably phosphorylates MITF-M. Glycogen synthase kinase 3b (GSK3b) is activated by cAMP signaling, and phosphorylates MITF-M at Ser-298, enhancing the DNA-binding ability of MITF-M. MITF-M activates transcription of the target genes, which are related to differentiation of melanocytes, such as tyrosinase family genes, and cell survival, such as Bcl2.

amino acid identity and 96% identity within the C-terminal acidic domain of 54 amino acids. CITED1 is a nonDNAbinding coactivator, and its C-terminal domain can function as a transcription activation domain through interacting with p300/CBP transcriptional coactivators (Fig. 13.4) (Yahata et al., 2000). CITED1 is a member of the CBP/p300-interacting transactivator with an ED-rich tail (CITED). CITED1 also interacts with Smad4 (Shioda et al., 1998) or estrogen receptors a and b (Yahata et al., 2001). Cited1 null mutant mice show growth restriction at 18.5 days post coitum, and most of them die shortly after birth, which suggest that Cited1 is required in trophoblasts for normal placental development and subsequently for embryo viability (Rodriguez et al., 2004). It has been shown that overexpression of CITED1 increases melanin in B16 melanoma (Nair et al., 2001).

BRN2 BRN2 (also called N-Oct3 or POU3F2) is a POU domaincontaining transcription factor, implicated in the development of the brain (reviewed by Nakai et al., 1995; Treacy and Rosenfeld, 1992). BRN2 mRNA was expressed in a range of human melanoma cell lines and was elevated compared with normal human melanocytes, whereas mRNA for Brn-2 was 248

A CANNTG motif is the consensus sequence of the binding site for a large family of transcription factors with a bHLH structure (Murre et al., 1989a, b). The bHLH structure is required for DNA binding and dimerization of transcription factors. A ubiquitous transcription factor, USF1, is one of such factors, containing a bHLH and a leucine zipper (LZ) structure (Gregor et al., 1990; Pognonec and Roeder, 1991). USF1 was initially identiﬁed as a nuclear protein that binds to the upstream element of the adenovirus major late promoter (Sawadogo and Roeder, 1985), and was then shown to be involved in the expression of various cellular genes (Carthew et al., 1987; Chodosh et al., 1987; Sato et al., 1990). USF1 was shown to bind in vitro to the mouse TRP-1 gene promoter (Yavuzer and Goding, 1994) and to TDE and TPE of the human tyrosinase gene (Yasumoto et al., 1994) using the antiserum against USF1. The binding of USF1 to TDE was also conﬁrmed using the USF1 produced by in vitro translation. In addition, many properties of the TDE-binding proteins are consistent with those of USF1, such as the elution proﬁle on ion-exchange chromatography and the binding patterns in the gel mobility shift assay (Yasumoto et al., 1994). Furthermore, coexpression of a USF1 cDNA remarkably increased the reporter gene expression under the control of the human tyrosinase promoter. USF1 has been identiﬁed as a target of the stress-responsive p38 kinase and could mediate UVinduced tyrosinase expression (Galibert et al., 2001). These results suggest that USF may be involved in efﬁcient transcription of the human tyrosinase gene. However, it should be noted that MITF protein does not form heterodimers with USF in vitro (Hemesath et al., 1994).

Tyrosinase Family Genes The tyrosinase family is directly involved in melanin pigment production and is speciﬁcally or preferentially expressed in pigment cells. Each family member has been extensively reviewed in other chapters; this chapter will focus on the transcriptional regulation of these genes. The tyrosinase family genes provide a good system to study the mechanisms of cell type-speciﬁc gene transcription. Such understanding is invaluable in studying how pigment cell differentiation is regulated. It is clear that multiple factors, including ubiquitous and cellspeciﬁc factors, must cooperate to direct pigment cell-speciﬁc transcription of the tyrosinase family genes. Here, we sum-

TRANSCRIPTIONAL REGULATION OF MELANOCYTE FUNCTION

marize some of the transacting factors acting on the tyrosinase family genes.

Tyrosinase or Albino (c) Locus Protein Tyrosinase (EC 1.14.18.1) is a rate-limiting enzyme in melanin biosynthesis that catalyzes the conversion of tyrosine to 3,4dihydroxyphenylalanine (DOPA), DOPA to DOPAquinone, and 5,6-dihydroxyindole to indole-5,6-quinone (Tripathi et al., 1992). The amino acid sequences of tyrosinase were deduced from chicken (Mochii et al., 1992), Japanese pond frog (Takase et al., 1992), Japanese quail, and snapping turtle (Yamamoto et al., 1992). They share signiﬁcant homology with mammalian tyrosinase. The 5¢ ﬂanking regions of the quail and turtle tyrosinase genes were cloned and compared with mammalian tyrosinase genes (Yamamoto et al., 1992). Through retrovirally mediated gene transfer, it was demonstrated that the quail and mouse promoters of about 500 bp are sufﬁcient to direct efﬁcient expression of a reporter mouse tyrosinase gene in cultured albino chick melanocytes but not in ﬁbroblasts and hepatocytes (Akiyama et al., 1994). This indicates that the quail promoter region used contains the cisacting element responsible for melanocyte-speciﬁc expression and that the mouse promoter is also functional in avian melanocytes. The quail promoter contains two copies of the element, containing the CANNTG motif, similar to the proximal melanocyte-speciﬁc element (p-MSE) of the mouse tyrosinase and TRP-1 genes (Yamamoto et al., 1992). These results suggest that a molecular mechanism for pigment cell-speciﬁc transcription of the tyrosinase gene is well conserved between birds and mammals. Mouse cDNA clones for tyrosinase have been isolated by several groups (Kwon et al., 1988; Müller et al., 1988; Terao et al., 1989; Yamamoto et al., 1987, 1989). The mouse tyrosinase gene, encoded at the albino (c) locus, was characterized and organized into ﬁve exons (Müller et al., 1988; Ruppert et al., 1988; Shibahara et al., 1990). The molecular basis of the BALB/c albinism was found as a point mutation, causing a Cys-to-Ser substitution at position 85 (codon 103) (Jackson et al., 1990; Shibahara et al., 1990; Yokoyama et al., 1990). Introduction of functional tyrosinase minigene constructs containing the 5¢ ﬂanking region into the albino mouse can rescue the albino phenotype (Beermann et al., 1990; Tanaka et al., 1992; Yokoyama et al., 1990). The promoter function of the mouse tyrosinase gene was analyzed in transgenic mice, showing that its 5¢ ﬂanking region of only 270 bp is sufﬁcient to provide cell type speciﬁcity (Klüppel et al., 1991) and developmental regulation in skin melanocytes and retinal pigment epithelium (Beermann et al., 1992). Within this 270-bp region, there are two copies of a CATGTG motif (positions –104 to –99 and –12 to –7). A high level of position-independent expression of a tyrosinase transgene has been achieved using a yeast artiﬁcial chromosome containing the 80-kb mouse tyrosinase gene and the 155-kb 5¢ ﬂanking region (Schedl et al., 1993), indicating that all cis-acting elements required for tyrosinase gene control are located in these DNA segments.

A DNase I hypersensitive site was identiﬁed about 15 kb upstream from the ﬁrst exon of the mouse tyrosinase gene (Porter et al., 1991). This hypersensitive site, located 12 kb upstream, was detected in mouse melanoma cells but not in mouse ﬁbroblasts (Ganss et al., 1994b). Functional analysis of the sequences corresponding to the DNase I hypersensitive site showed the property of a strong cell-speciﬁc enhancer in transgenic mice as well as in cultured cells (Ganss et al., 1994b). It was also shown that full enhancer activity in transient expression assays is detected with a minimal sequence of 200 bp located at –12.1 kb. Two protein-binding regions, hs1 and hs2, were identiﬁed within the 200-bp enhancer region by DNase I footprinting analysis. Interestingly, either the hs1 or the hs2 region contains a copy of the CANNTG motif. However, by gel retardation assays, the protein-binding activity was detected only with hs1 oligonucleotide, but not with hs2 oligonucleotide, suggesting that protein binding to hs1 is a prerequisite for the binding factors to hs2 (Ganss et al., 1994b). A palindromic sequence 5¢-TGACTTTGTCA-3¢ is present in the hs1 region and is similar to the binding motif of CREB (cAMP-responsive element binding) transcription factor and an AP1 (activator protein 1) binding site, which is recognized by members of the fos/jun family of transcription factors. The enhancer function of the DNase I hypersensitive site of the mouse tyrosinase gene was also conﬁrmed in transgenic mice by other investigators (Porter and Meyer, 1994). They also suggested that the distinct regulatory elements are required for the expression of tyrosinase gene in neural crestderived melanocytes and RPE. The 270-bp upstream region of the mouse tyrosinase gene was analyzed in detail (Ganss et al., 1994a), because this region is sufﬁcient to confer cell-speciﬁc and developmentally regulated expression of the tyrosinase gene in the transgenic mice (Beermann et al., 1992). Within this 270-bp region, two positive regulatory elements (positions –245 to –230 and –104 to –93) and one negative regulatory element (position –195 to –125) were identiﬁed by transient expression assays (Ganss et al., 1994a). The positive regulatory element, located between –104 and –93, coincides with the M-box, the 11-bp cis-acting element containing the CATGTG motif of the mouse TRP-1 gene (Lowings et al., 1992). It was also shown that MITF activates the mouse tyrosinase gene promoter (Yasumoto et al., 1995). However, the functional analysis of the two positive elements in transgenic mice indicates that these two elements do not determine pigment production in vivo but rather modulate transcription of the mouse tyrosinase gene (Ganss et al., 1994c). Kwon et al. (1987a) isolated a portion of human tyrosinase cDNA by screening a melanocyte expression cDNA library with antityrosinase polyclonal antibodies. Subsequently, the entire structure of human tyrosinase was deduced from the full-length cDNAs (Bouchard et al., 1989; Shibahara et al., 1988; Takeda et al., 1989). The catalytic function of a cloned tyrosinase cDNA was assessed by transient transfection assays, revealing that tyrosinase possesses the two catalytic activities: tyrosine hydroxylase and dopa oxidase (Takeda 249

CHAPTER 13

et al., 1989). The human tyrosinase gene was mapped to chromosome 11q14–q21, and a second “tyrosinase-related” sequence mapped to 11p11.2–cen (Barton et al., 1988). Tomita et al. (1989) clariﬁed the molecular basis of oculocutaneous albinism (type I) by ﬁnding a single base insertion in the exon sequence of the patient’s tyrosinase gene. This insertion causes a frameshift and introduces a premature termination codon. The patient has the mutant gene at both alleles, and the truncated protein encoded by the mutant gene is not functional. The human tyrosinase gene is organized into ﬁve exons (Tomita et al., 1989), and its size is greater than 70 kb (Takeda et al., 1990). The nucleotide sequence of the 5¢ ﬂanking region of the human tyrosinase gene was also reported (Kikuchi et al., 1989). The promoter function of the mutant gene is conﬁrmed by in vitro transcription using melanoma whole cell extracts (Takeda et al., 1990). The structural organization and the nucleotide sequence of the 5¢ ﬂanking region of the human tyrosinase gene were also reported (Giebel et al., 1991; Ponnazhagan et al., 1994). It was found that the second site, mapped to chromosome 11p11.2–cen by Barton et al. (1988), represents the truncated human tyrosinase pseudogene, which consists of the nucleotide sequence almost identical to that of exons 4 and 5 (Giebel et al., 1991; Takeda et al., 1991). Shibata et al. (1992a) identiﬁed the 39-bp enhancer element of the human tyrosinase gene about 1.8 kb upstream from the transcription initiation site, which is responsible for pigment cell-speciﬁc expression of a reporter gene. The enhancer element was narrowed to the 20-bp sequence, termed tyrosinase distal element (TDE) (positions –1861 to –1842) (Yasumoto et al., 1994). TDE, containing a CATGTG motif in its center, was speciﬁcally bound by MITF-M (Yasumoto et al., 1994, 1997). There are two additional copies of the CATGTG motif in the human tyrosinase promoter region (positions –104 to –99 and –12 to –7) (Yasumoto et al., 1994). The 20-bp region (positions –112 to –93), containing the CATGTG motif (positions –104 to –99), was identiﬁed as a weak pigment cell-speciﬁc promoter, termed tyrosinase proximal element (TPE) (Yasumoto et al., 1994). It is noteworthy that the 1.0-kb 5¢ ﬂanking region of the human tyrosinase gene containing TPE but lacking TDE was sufﬁcient to confer cellspeciﬁc expression of the tyrosinase gene in transgenic mice (Tanaka et al., 1992). However, no pigmented melanocytes were detected in the epidermis of these mice. It is therefore conceivable that TDE is required for efﬁcient and correct expression of the human tyrosinase gene in a pigment cellspeciﬁc manner. Suzuki et al. (1994) have reported that overexpression of neuroﬁbromin cDNA in MeWo melanoma cells, which are deﬁcient in neuroﬁbromin, enhanced the expression of a reporter gene under the control of the tyrosinase promoter. Neuroﬁbromin is responsible for neuroﬁbromatosis type 1 (NF1) or von Recklinghausen’s disease, which is a common autosomal dominant disorder characterized by the presence of neuroﬁbromas, café-au-lait spots, Lisch nodules (hamartomas in the iris), bone deformity, and an increased frequency of 250

malignancies (Cawthon et al., 1990; Viskochil et al., 1990; Wallace et al., 1990). Neuroﬁbromin has a domain related to mammalian ras GTPase-activating protein (GAP) (Xu et al., 1990). Notably, the GAP-related domain of neuroﬁbromin is mainly responsible for the activation of the tyrosinase promoter (Suzuki et al., 1994). Likewise, neuroﬁbromin has been shown to activate the promoter activity of the human DCT gene, as judged by transient cotransfection assays (Suzuki et al., 1998). Neuroﬁbromin may modulate intracellular signaling, which in turn stimulates the activity of a factor required for melanocyte-speciﬁc transcription of the tyrosinase and DCT genes.

Tyrosinase-related Protein 1 A pigment cell-speciﬁc cDNA, pMT4, was cloned from a B16 mouse melanoma cDNA library by differential hybridization (Shibahara et al., 1986). The expression of pMT4 mRNA is restricted to the skin and pigmented melanoma cells. The deduced structure of the pMT4-encoded protein shares signiﬁcant homology with Neurospora tyrosinase and contains the several domains thought to be characteristic of tyrosinase, such as a signal sequence, two potential Cu-binding regions, a transmembrane domain, and potential glycosylation sites. Furthermore, transient expression of pMT4 revealed that its protein product is reactive with monoclonal antibodies TMH1 and TMH-2, which have been reported as antityrosinase monoclonal antibodies (Tomita et al., 1985). The pMT4 protein was therefore reported as tyrosinase (Shibahara et al., 1986). However, when its gene was mapped to the brown (b) locus on mouse chromosome 4, which determines the type of melanin produced, it was shown to be different from tyrosinase (Jackson, 1988). The pMT4 protein is now known as TRP-1, and monoclonal antibodies TMH-1 and TMH-2 have been established as anti-TRP-1 antibodies (Tomita et al., 1991). The function of TRP-1 was identiﬁed as DHICAoxidase, catalyzing the conversion of DHICA to indole– quinone–carboxylic acid (Jiménez-Cervantes et al., 1994; Kobayashi et al., 1994). Thus, TRP-1 is the ﬁrst gene of the tyrosinase gene family to be cloned, but its exact function was established last. TRP-1 or brown (b) locus protein is composed of 537 amino acids, including a signal peptide with a molecular mass of 58 kDa (Shibahara et al., 1986). Sequence analysis of the BALB/c mouse TRP1 gene revealed ﬁve base differences in the exon regions compared with the sequence of the C57BL/6 gene (Shibahara et al., 1991). One is a deletion of three nucleotides in the 5¢-untranslated region, and the other four are point mutations within the protein coding region: two missense mutations and two silent mutations (Zdarsky et al., 1990). These two missense mutations are also found in another b-mutant mouse DBA/2 (Shibahara et al., 1992). The original b mutation has been identiﬁed as a Cys-to-Tyr substitution at position 86 (codon 110) (Jackson et al., 1990; Zdarsky et al., 1990). The mutant TRP-1 containing Tyr-110 is not reactive with an anti-TRP-1 monoclonal antibody TMH-1 (Shibahara et al., 1992).

TRANSCRIPTIONAL REGULATION OF MELANOCYTE FUNCTION

The mouse TRP1 gene is about 15~18 kb long and organized into eight exons (Jackson et al., 1991; Shibahara et al., 1991). Jackson et al. (1991) assigned several potential transcriptional initiation sites, and one of them is the same as that assigned by Shibahara et al. (1991). In this review, we have used the numbering that starts with the upstream initiation site (position 1) (Jackson et al., 1991); thus, the initiation site assigned by Shibahara et al. (1991) is the nucleotide at position 5. The promoter function of the mouse TRP1 gene was analyzed using an in vitro transcription system, suggesting that the DNA segment (position –34 to +158) is sufﬁcient to direct pigment cell-speciﬁc transcription in melanoma whole cell extracts but not in HeLa whole cell extracts (Shibahara et al., 1991). On the other hand, through transient transfection assays, Lowings et al. (1992) have identiﬁed the regulatory region consisting of both positive and negative elements, which confers efﬁcient expression on a reporter gene in pigmented melanoma cells but not in nonmelanoma cells. The positive element contains the nonconsensus octamer motif, 5¢ATTTGAAT-3¢, and has been shown to be bound in vitro by the ubiquitous OCT-1 transcription factor. In addition, a minimal promoter extending between –44 and +107 is sufﬁcient for cell type-speciﬁc expression, and it contains the 11bp cis-acting element, termed the M-box. The M-box contains the CATGTG motif and is required for a basal level of expression of a reporter gene in B16 mouse melanoma cells (Lowings et al., 1992). Yasumoto et al. (1995) have found through transient expression analysis that MITF transactivates the mouse TRP-1 promoter, mainly via the M-box. The structure of human TRP-1 has been deduced from its cDNA sequence (Chintamaneni et al., 1991; Cohen et al., 1990), sharing about 93% amino acid identity with a mouse counterpart. Human TRP-1 is composed of 527 amino acids and is shorter than mouse TRP-1 by 10 amino acids at the carboxyl-terminus (Cohen et al., 1990). Human TRP-1 is also known as a melanoma antigen gp75, which is potentially autoimmunogenic, because immunoglobulin G (IgG) antibodies against gp75 are detected in a patient with metastatic melanoma (Vijayasaradhi et al., 1990). Subsequently, an informative single base change was found in the full-length human TRP-1 cDNA, pHT2a, that was isolated from a MeWo human melanoma cDNA library (Takimoto et al., 1995); namely, the HindIII site (AAGCTT) present in the reported sequence (Cohen et al., 1990) is disrupted in pHT2a because of the Cto-T transition at position 750. The human TRP-1 gene contains the C residue at position 750, favoring the sequence previously reported, which was derived from a different human melanoma cell line (Cohen et al., 1990). This base change is not associated with an amino acid substitution and may represent polymorphism. The human TRP1 gene has been mapped to chromosome 9p22–pter (Chintamaneni et al., 1991), and the 5¢ ﬂanking region and exon 1 of the TRP1 gene have been characterized and the promoter function assessed by transient expression assays (Shibata et al., 1992b). The structural organization of

the human TRP1 gene has been characterized (Sturm et al., 1995). Unlike the mouse TRP1 gene (Yasumoto et al., 1995), the pigment cell-speciﬁc promoter function of the human TRP1 gene was not detected by transient expression assays in cultured cells with the 5¢ ﬂanking region of up to 3.5 kb (Shibata et al., 1992b; Yasumoto et al., 1997; Yokoyama et al., 1994b).

DOPAchrome Tautomerase/Tyrosinase-related Protein 2 The phenotype of homozygous slaty mice includes the dilution of coat color and premature hair loss. It was proposed that DCT functions to prevent the accumulation of dihydroxyindole, a more toxic product of DOPAchrome generated spontaneously (Jackson et al., 1992). The DCT gene was mapped to the slaty locus on mouse chromosome 14 and is mutated at the slaty locus, which is an arginine to glutamine change at the ﬁrst copper binding site (Jackson et al., 1992). It was then conﬁrmed by transient expression assays of cDNA that mouse TRP-2 possesses the catalytic activity of DOPAchrome tautomerase and the slaty mutation dramatically decreases the catalytic function (Kroumpouzos et al., 1994). The mouse DCT gene is expressed in migratory melanoblasts shortly after they left the neural crest (Steel et al., 1992); expression of the DCT gene precedes that of the tyrosinase and TRP1 genes. Likewise, DCT expression is detected in the developing telencephalon and the endolymphatic duct, in which expression of the tyrosinase and TRP1 genes was undetectable (Steel et al., 1992). Furthermore, DCT mRNA is expressed in a glioblastoma (Suzuki et al., 1998) and the retinoblastoma (Udono et al., 2001). Thus, the regulation of DCT gene expression is different from that of tyrosinase and TRP1. The structure of the mouse DCT gene has been reported (Budd and Jackson, 1995), and its promoter region contains the M-box. Human DCT shares about 40% amino acid identity with human tyrosinase or TRP-1 (Bouchard et al., 1994; Cassady and Sturm, 1994; Yokoyama et al., 1994a). It has also been conﬁrmed that DCT possesses the catalytic activity of DOPAchrome tautomerase by transient expression assays of a human cDNA (Yokoyama et al., 1994a). The human DCT gene has been mapped to chromosome 13q32 (Bouchard et al., 1994). The 5¢ ﬂanking region and the portion of the human DCT gene have been cloned and characterized (Sturm et al., 1995; Yokoyama et al., 1994b). Comparison of the nucleotide sequence with that of the other tyrosinase gene family members reveals two putative cis-acting elements (–138/–128 and –34/–21), which are similar to those of the M-box (Lowings et al., 1992) and a pigment cell-speciﬁc promoter (Shibahara et al., 1991) respectively. The M-box of the human DCT gene (–138 to –128) is identical to the equivalent element of the mouse DCT gene. Transient expression assays revealed that the 5¢ ﬂanking region of the human DCT gene upstream from a luciferase 251

CHAPTER 13

reporter gene is more efﬁciently expressed in MeWo melanoma cells than in HeLa cells (Yokoyama et al., 1994b), indicating the presence of the pigment cell-speciﬁc regulatory element(s). It was then found that the 32-bp regulatory element responsible for pigment cell-speciﬁc expression is located between –447 and –415, which was termed the DCT distal enhancer (Amae et al., 2000) (Fig. 13.3), although this enhancer alone is not sufﬁcient to confer cell type-speciﬁc expression. An additional element, located between –268 and –56, is required for basal transcription, but again this element by itself is not sufﬁcient to confer pigment cell-speciﬁc expression. The latter DNA segment (–268/–56) was tentatively termed the proximal region. Notably, it has been shown that the 71-bp region (–415/–345), immediately downstream from the DCT distal enhancer, is required for activation of the DCT promoter by neuroﬁbromin (Suzuki et al., 1998). Thus, multiple regulatory elements are required for the pigment cell-speciﬁc transcription of the DCT gene. Yasumoto et al. (1997) have reported that MITF is unable to trans-activate the DCT promoter in transient transfection assays, despite the fact that MITF binds in vitro to the DCT M-box. Subsequently, it has been conﬁrmed that the M-box of the DCT promoter is bound by MITF-M in vivo (Takeda et al., 2003). Furthermore, an experimentally truncated MITF protein lacking the carboxyl-terminal 125 amino acids activated the tyrosinase promoter less efﬁciently than did MITF but, surprisingly, the truncated MITF activated the DCT promoter (Yasumoto et al., 1997). These results suggest that the carboxyl-terminus of MITF contains a transcriptional activation domain that is also involved in deﬁning the binding sites for MITF. The synergism between LEF-1 and MITF-M is responsible for the transcriptional regulation of the DCT gene but through a different mechanism from that of the M promoter (Saito et al., 2002; Yasumoto et al., 2002). DCT may be important for the survival of melanocytes. The ﬁnding that the vit mutation impairs the synergism with LEF-1 on the DCT promoter but not on the M promoter is of particular interest in view of the phenotype of homozygous Mitf vitiligo (Mitf vit) mice that appear normal as young with uniformly lighter color but show aging-dependent melanocyte loss (Lerner et al., 1986). The Asp222Asn substitution in the helix 1 of Mitf-M represents a molecular lesion of the recessive Mitf vit (Steingrímsson et al., 1994), and the Mitf-Mvit protein is able to bind in vitro to DNA (Hemesath et al., 1994). Thus, the Mitf vit mutation does not profoundly alter the fetal development of melanocytes but impairs the postnatal maintenance of follicular melanocytes, which could be explained by the differential effects of the Mitf vit mutation on the synergism with LEF-1 on the M promoter and the DCT promoter (Saito et al., 2002).

Pmel-17 Protein or Silver Locus Protein A portion of cDNA coding for Pmel-17 protein was initially cloned by screening a human melanocyte cDNA expression library with antityrosinase polyclonal antibodies (Kwon et al., 1987b). A full-length Pmel-17 cDNA was subsequently iso252

lated, and Pmel-17 protein is composed of 645 amino acids with a molecular mass of 68.6 kDa (Kwon et al., 1991). Pmel17 protein contains a putative leader sequence, a potential membrane anchor segment, and relatively high numbers of serine and threonine residues. There are ﬁve restricted regions of amino acid similarity among tyrosinase, TRP-1, and Pmel17. The Pmel-17 gene has been mapped to human chromosome 12pter–q21, and its mouse homolog has been mapped to the distal region of mouse chromosome 10, a region known to carry the coat color locus silver (si) (Kwon et al., 1991). The recessive silver mutation causes progressive graying of hair due to the loss of functional follicular melanocytes. The silver gene, a mouse homolog of Pmel-17, is thought to act at the subcellular level and is preferentially expressed in melanocytes (Kwon et al., 1987b). Kwon et al. (1995) found that the mouse silver mutation is caused by a single base insertion in the putative cytoplasmic domain of the mouse homolog of Pmel-17 protein. Pmel-17 is expressed in RPE, because the two cDNA clones derived from chicken and bovine RPE turned out to be a homolog of Pmel-17. The former cDNA encodes a melanosomal matrix protein MMP115 of chicken RPE (Mochii et al., 1991), and the latter encodes the bovine retinal pigment epithelial protein RPE-1 (Kim and Wistow, 1992). In particular, MMP115 was shown to be a useful marker for studying the molecular mechanisms for the differentiation of RPE, because the MMP115 gene was transcriptionally inactivated during the process of dedifferentiation of RPE to a bipotent cell, which can be committed to either the lens cell or RPE (Mochii et al., 1988). Furthermore, expression of Pmel-17 mRNA was remarkably increased following the treatment of murine or human melanoma cells with b-MSH or 3-isobutyl1-methylxanthine (Kwon et al., 1987b), and thus the regulation of its gene expression is of interest.

Transcription Factors Acting at Retinal Pigment Epithelium Correct speciﬁcation and differentiation of RPE underlie eye morphogenesis, and the impairment of RPE development could lead to severe malformation of the eye, such as anophthalmia (Hero et al., 1990). RPE is derived from the optic cup of embryonic brain and differentiates into a single layer of cells interposed between the neural retina and the vascular-rich choroids, thereby constituting the blood–retinal barrier. At the apical side of RPE, many villi interdigitate with the outer segments of photoreceptors, by which RPE is responsible for phagocytosis of outer segments of photoreceptors (Bok, 1988) and for uptake, processing, and transport of retinoids that are essential for visual function (Bok, 1993). RPE therefore plays important roles in the survival and function of photoreceptors in the adult eye. The mutant homozygous mice for Mitf exhibit depigmentation and hyperproliferation of the RPE and the formation of an ectopic neuroretina at the expense of the dorsal RPE, resulting in microphthalmia (Silver, 1979). In the homozygous Mitf VGA-9 mice, where Mitf is not expressed, expression of

TRANSCRIPTIONAL REGULATION OF MELANOCYTE FUNCTION

tyrosinase and TRP1 genes is reduced in RPE, suggesting that MITF is required for efﬁcient transcription of the tyrosinase and TRP1 genes in RPE as well as in melanocytes (Nakayama et al., 1998). But Mitf is dispensable for Dct expression, as expression of Dct is not affected in Mitf VGA-9 mice (Nakayama et al., 1998). MITF-A is preferentially expressed in a human RPE line of fetal origin and is also expressed in many cell types (Amae et al., 1998; Fuse et al., 1999; Udono et al., 2000). Domain A of MITF-A shares signiﬁcant amino acid identity with the N-terminus of TFE3 (Rehli et al., 1999; Yasumoto et al., 1998). In addition, three consecutive portions, covering the entire domain A, are aligned to the equivalent portions of cytoplasmic retinoic acid-binding protein (CRABP) (Shibahara et al., 2001). Such similarity is of interest, in view of the phenotype of a recessive Mitf mutant, Mitf vit (Lerner et al., 1986; Steingrímsson et al., 1994), which shows late-onset retinal degeneration and abnormalities in retinoid metabolism (Smith et al., 1994). It is therefore conceivable that the Asp-to-Asn substitution in the helix 1 of MITF-A or other MITF isoforms may impair the interaction with a hitherto unidentiﬁed protein in RPE. Transcription of the DCT gene is activated by OTX2 (Takeda et al., 2003). OTX2 is a homolog of the Drosophila homeobox-containing gene orthodenticle (otd), a member of a highly conserved family of homeodomain-containing transcription factors, and is expressed not only at the earliest stage of eye morphogenesis including prospective RPE (MartinezMorales et al., 2001; Simeone et al., 1992, 1993), but also in RPE of the postnatal and adult mouse eye (Baas et al., 2000). In addition, Otx2 null homozygous mice exhibit the absence of the forebrain and embryonic lethality (Acampora et al., 1995), whereas the heterozygotes survive until birth but exhibit anomalies in RPE and retinal development (Matsuo et al., 1995). OTX2 interacts with MITF and synergistically activates tyrosinase and TRP1 genes in RPE (MartinezMorales et al., 2003). It has also been reported that MITF interacts with PAX6 in RPE (Planque et al., 2001). PAX6 is a member of the pairedhomeodomain family of transcription factors (Walther and Gruss, 1991), mutations in which cause the aniridia syndrome in humans (Glaser et al., 1992) and the small eye phenotype in mice (Hill et al., 1991). The interaction with PAX6 and MITF abolishes the DNA binding of MITF, leading to a reduction in MITF-mediated activation in RPE (Planque et al., 2001). Several RPE-speciﬁc proteins were identiﬁed, including tyrosinase family genes, and cDNAs coding for some of them were cloned. A typical example of such proteins is Pmel-17, also known as MMP115 or RPE-1, which was initially cloned as an RPE-speciﬁc gene (Kim and Wistow, 1992; Mochii et al., 1991). Another example is RPE65, which was cloned as a novel RPE-speciﬁc microsomal protein with a tissue-speciﬁc monoclonal antibody RPE9 (Hamel et al., 1993). Bovine RPE65 consists of 533 amino acid residues with no predicted transmembrane domains. RPE65 mRNA was detected in RPE

but not in other ocular tissues, suggesting that RPE65 is an RPE-speciﬁc protein. The human RPE65 gene has been mapped to chromosome 1p31 (Hamel et al., 1994). The gene for ocular albinism type 1 (OA1), located at chromosome Xp22.3–22.2, was identiﬁed by positional cloning (Bassi et al., 1995). OA1 is an X-linked disorder characterized by severe impairment of visual acuity, retinal hypopigmentation, and the presence of macromelanosomes. The OA1 gene encodes a protein of 424 amino acids with several putative transmembrane domains and is preferentially expressed in retina as well as in melanoma. Five intragenic deletions and a CG dinucleotide insertion causing a premature stop codon were identiﬁed in unrelated OA1 patients. It is of interest to study the pathogenesis of OA1, because the OA1 gene is not RPE speciﬁc and is also expressed in melanoma, like many other pigment-related genes. The tyrosinase family genes therefore provide a good system to study the mechanism responsible for RPE-speciﬁc gene expression, which in turn allows us to compare regulation in melanocytes.

Perspectives The MITF gene consists of widely spaced multiple promoters, which generate not only the diversity in the transcriptional regulation of these promoters but also the structurally different isoforms. The MITF gene provides a good model for studying the mechanism of promoter selectivity during development or under certain metabolic conditions. The isoform multiplicity of MITF also provides functional diversity and redundancy. Future analysis of MITF isoforms in vivo is required to assess the role of each MITF isoform in differentiation and development of the pigment cell and other cell types. It is also important to identify the interacting partners with the unique N-terminal domains as well as the bHLH-LZ domain that modulate the function of MITF isoforms. Such studies will increase our understanding of the regulation of development and differentiation of the pigment cell. Development of gene therapy and chemotherapy toward melanoma are deﬁnitely major goals of pigment cell research. Retinal degeneration of diverse etiologies, either acquired or inherited, accounts for a major cause of aging-dependent visual impairment and blindness in the developed world. It is therefore of signiﬁcance to understand the mechanism by which various transcription factors maintain the survival or function of RPE in the adult retina. Furthermore, the elucidation of the difference in transcriptional regulation of the tyrosinase family genes between RPE and melanocytes is of particular signiﬁcance. It is also challenging to explore the possibility of cell therapy targeted for RPE in order to deliver a healthy RPE into an impaired retina.

Acknowledgments We thank Dr Hiroaki Yamamoto (Tohoku University) for 253

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helpful discussions. We also thank Ken-ichi Yasumoto, Kazuhiro Takahashi, and other colleagues who contributed to the work on pigment cells.

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