Molecular Basis Of Epithelial Appendage Morphogenesis (Molecular Biology Intelligence Unit)

M OLECULAR BIOLOGY I N T E L L I G E N C E U N I T

1

Cheng-Ming Chuong

Molecular Basis of Epithelial Appendage Morphogenesis

R.G. LANDES C O M P A N Y

MOLECULAR BIOLOGY INTELLIGENCE UNIT 1

Molecular Basis of Epithelial Appendage Morphogenesis Cheng-Ming Chuong Department of Pathology University of Southern California Los Angeles, California, U.S.A.

R.G. LANDES COMPANY AUSTIN, TEXAS U.S.A.

MOLECULAR BIOLOGY INTELLIGENCE UNIT Molecular Basis of Epithelial Appendage Morphogenesis R.G. LANDES COMPANY Austin, Texas, U.S.A. Copyright © 1998 R.G. Landes Company All rights reserved. No part of this book may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. Printed in the U.S.A. Please address all inquiries to the Publishers: R.G. Landes Company, 810 South Church Street, Georgetown, Texas, U.S.A. 78626 Phone: 512/ 863 7762; FAX: 512/ 863 0081

ISBN: 1-57059-490-2

While the authors, editors and publisher believe that drug selection and dosage and the specifications and usage of equipment and devices, as set forth in this book, are in accord with current recommendations and practice at the time of publication, they make no warranty, expressed or implied, with respect to material described in this book. In view of the ongoing research, equipment development, changes in governmental regulations and the rapid accumulation of information relating to the biomedical sciences, the reader is urged to carefully review and evaluate the information provided herein.

Library of Congress Cataloging-in-Publication Data

CIP applied for but not yet received at time of publication

MOLECULAR BIOLOGY INTELLIGENCE UNIT 1 PUBLISHER’S NOTE

Molecular Basis of Epithelial Appendage Morphogenesis

Landes Bioscience produces books in six Intelligence Unit series: Medical, Molecular Biology, Neuroscience, Tissue Engineering, Biotechnology and Environmental. The authors of our books are acknowledged leaders in their fields. Topics are unique; almost without exception, no similar books exist on these topics. Our goal is to publish books in important and rapidly changing areas of bioscience for sophisticated researchers and clinicians. To achieve this goal, we have accelerated our publishing program to conform to the fast pace at which information grows in bioscience. Most of our books Department are published of within 90 to 120 days of receipt of Pathology the manuscript. We would like to thank our readers for their University of Southern California continuing interest and welcome any comments or suggestions they Los Angeles, California, U.S.A. may have for future books.

Cheng-Ming Chuong

Judith Kemper Production Manager R.G. Landes Company

R.G. LANDES COMPANY AUSTIN, TEXAS U.S.A.

CONTENTS Part I: Overview 1. Morphogenesis of Epithelial Appendages: Variations on Top of a Common Theme and Implications in Regeneration ....................... 3 Cheng-Ming Chuong The Epithelial Appendage Paradigm ...................................................... 3 Studies on Epithelial Appendage Morphogenesis ................................. 5 What Is in this Book? .............................................................................. 8 Organogenesis, Histogenesis, and New Growth .................................. 10 2. Ectodermal Dysplasia: A Synthesis Between Evolutionary, Developmental, and Molecular Biology and Human Clinical Genetics ................................................................. 15 Harold C. Slavkin, Lillian Shum and Glen H. Nuckolls Introduction ........................................................................................... 15 Ectodermal Dysplasias ........................................................................... 17 Anhydrotic Ectodermal Dysplasia (EDA) ............................................ 18 Hierarchical Regulation and Feedback Loops in Epithelial-Mesenchymal Interactions .......................................... 23 Animal Models for Ectodermal Dysplasias .......................................... 28 Prospectus .............................................................................................. 30 3. Why Study Hair Follicles? A Personal Account ..................................... 39 Margaret Hardy Fallding Part II: Morphogenesis of Epithelial Appendages on the Body Surface 4. Variations of Cutaneous Appendages: Regional Specification and Cross-Species Signals ................................. 45 Danielle Dhouailly, Fabrice Prin, Benoit Kanzler and Jean P. Viallet Introduction ........................................................................................... 45 The Formation of Embryonic Skin Varies According to the Different Body Regions ........................................ 46 The Predermal Cells Are Regionally Specifically Determined but Need a Systemic Nonspecific FGF-Like Epidermal Message to Acquire Their Dermal Inductive Potentialities ........................... 46 Regional-Specific Dermal Messages Which Control the First Step of Cutaneous Appendage Formation Can Be Correctly Interpreted by an Epidermis from a Different Class ....................... 47 The Epidermis Does Not Possess Equivalent Competence All Over the Body .............................................................................. 49 Molecular Specification Along the Anteroposterior and Dorsoventral Axes Act Together Toward Skin Regionalization ..... 50 Conclusion ............................................................................................. 53

5. Feather Morphogenesis: A Model of the Formation of Epithelial Appendages ........................................................................ 57 Cheng-Ming Chuong and Randall B. Widelitz Introduction........................................................................................... 57 Overview of Feather Morphogenesis .................................................... 62 Macropatterning: Setting Up the Tract Fields ..................................... 64 Micropatterning: Generation of Feather Primordia Within the Tract ................................................................................ 67 Anterior-Posterior Axis: Orientation ................................................... 69 Proximal-Distal Axis: Elongation ......................................................... 70 Feather Filament Morphogenesis ......................................................... 71 Perspectives ............................................................................................ 72 6. Principles of Hair Follicle Morphogenesis............................................. 75 Michael Philpott and Ralf Paus Introduction........................................................................................... 75 Morphology of Hair Follicle Development .......................................... 82 Principles of Hair Follicle Induction. ................................................... 86 Signaling Events in Hair Follicle Development ................................... 86 A Working Model: Molecular Controls of Hair Morphogenesis ..... 103 7. Growth of the Hair Follicle: A Cycling and Regenerating Biological System .................................................... 111 Kurt Stenn, S. Parimoo and Steve Prouty Introduction......................................................................................... 111 Antler Formation ................................................................................. 113 Cycle of the Hair Follicle ..................................................................... 114 Hair Follicle Growth Controls—A Mechanistic Overview ............... 122 Conclusions ......................................................................................... 124 8. Development of Melanocytes from Neural Crest Progenitors ........... 131 Laure Lecoin, Ronit Lahav, Elisabeth Dupin and Nicole Le Douarin Introduction......................................................................................... 131 Early Migration and Commitment of Pigment Cells ........................ 131 From Genes to Mechanisms Regulating Pigmentation ..................... 138 Concluding Remarks ........................................................................... 147 Part III: Morphogenesis of Epithelial Appendages Within the Body 9. The Teeth as Models for Studies on the Molecular Basis of the Development and Evolution of Organs ..................................... 157 Irma Thesleff and Johanna Pispa Introduction......................................................................................... 157 Evolution of Teeth ............................................................................... 158 The Neural Crest ................................................................................. 160

Patterning of Teeth .............................................................................. 161 Morphogenesis of Individual Teeth: Regulation by Epithelial-Mesenchymal Interactions .................... 162 The Enamel Knot as a Signaling Center ............................................. 166 Signaling Networks and Transcription Factors ................................. 169 Disrupted Tooth Development .......................................................... 171 Concluding Remarks ........................................................................... 173 10. Epithelial-Mesenchymal Interactions in Gastrointestinal Development ......................................................... 181 Drucilla J. Roberts Introduction ......................................................................................... 181 Initial Development of the Vertebrate Lumenal Gut: Inductive Events .............................................................................. 182 Patterning and Epithelial Differentiation: Inductive Events ............. 184 Genetic Controls of Gut Development .............................................. 186 Genetics of Gut Development in Drosophila .................................... 187 Genetic Controls of Vertebrate Gut Pattern ...................................... 189 11. Endodermal Appendage Formation: Morphogenetic Mapping of Dorso-Ventral Patterning of the Anterior Foregut and Development of Lung and Thyroid Primordia ............................ 203 Parviz Minoo, Shioko Kimura and Robert deLemos Introduction ......................................................................................... 203 Progenitor Field Specification ............................................................ 203 Pattern Formation in the Anterior Foregut ....................................... 205 Further Regionalization ...................................................................... 206 Possible Functions for Nkx-2.1 ........................................................... 209 A Simple Molecular Morphogenetic Map for Lung and Thyroid Development .............................................................. 210 12. Lung Epithelial Morphogenesis: Integrated Functions of Transcriptional Factors, Peptide Growth Factors, Extracellular Matrix, Physiological and Environmental Factors ....... 215 David Warburton, Guillermo Flores-Delgado, Ding Bu, Kathryn D. Anderson and Richard E. Olver Introduction ......................................................................................... 215 Lessons on Branching Morphogenesis from Drosophila Genetics ............................................................... 220 Molecular Determinants of Lung Morphogenesis ............................. 221 Physiological Determinants of Lung Morphogenesis ........................ 229 Perspectives on Lung Regeneration .................................................... 230

Part IV: Molecular Mechanism 13. Early Molecular Events in Feather Morphogenesis: Induction and Dermal Condensation .................................................. 243 Randall B. Widelitz and Cheng-Ming Chuong Introduction......................................................................................... 243 Molecules in the Feather Inductive Phase .......................................... 244 Molecules in the Dermal Condensation Phase .................................. 251 Size Determination: Boundary and Spacing ...................................... 255 Expression Modes and Molecular Cascades ...................................... 256 Summary .............................................................................................. 256 14. Signaling Loops in the Reciprocal Epithelial-Mesenchymal Interactions of Mammalian Tooth Development ............................... 265 Yi-Ping Chen and Richard Maas Introduction......................................................................................... 265 General Models for Signaling Loops in Tissue Interactions ............. 267 Expression of Growth Factors Is Dynamic During Early Tooth Development .................................................. 267 Expression of Transcription Factors Is Associated with Inductive Processes ................................................................. 271 Growth Factors as Signals Mediating Reciprocal Epithelial-Mesenchymal Interactions ............................................ 273 MSX Genes Regulate the Expression of Inductive Signals ................ 275 15. Topobiology of the Hair Follicle: Adhesion Molecules as Morphoregulatory Signals During Hair Follicle Morphogenesis ...... 283 Sven Müller-Röver and Ralf Paus Introduction......................................................................................... 283 Principles of Topobiology in Morphogenesis .................................... 283 Topobiological Questions in Hair Follicle Morphogenesis .............. 291 Perspectives .......................................................................................... 302 16. Late Events and the Regulation of Keratinocyte Differentiation in Hair and Feather Follicles ................................................................. 315 George E. Rogers, Stephanie Dunn and Barry Powell Introduction......................................................................................... 317 Growth and Late Biochemical Events in the Hair Follicle ................ 323 The Genes for Structural Proteins and Their Expression in the Cortex and Cuticle During Hair Growth ............................ 324 Comparison of Hair Keratinization with Avian Keratin Formation in the Feather Follicle ...................................................................... 331 Concluding Remarks ........................................................................... 332

Part V: Models 17. Epithelial Morphogenesis: A Physico-Evolutionary Interpretation ............................................... 341 Stuart A. Newman Introduction ......................................................................................... 341 The Earliest Metazoa: Excitable Soft Matter ...................................... 343 Consequences of Differential Adhesion in Fluid Epithelia ............... 344 Pattern Formation in Excitable Epithelia ........................................... 347 Segmentation ....................................................................................... 348 Shape Change in Epithelial Sheets ...................................................... 352 Conclusion ........................................................................................... 354 18. Periodic Pattern Formation of the Feathers ........................................ 359 Han-Sung Jung, Cheng-Ming Chuong Introduction ......................................................................................... 359 Surveys of Existing Models ................................................................. 360 A Novel Integrated Model of Feather Pattern Formation ................. 364 Conclusions ......................................................................................... 366 19. Gene Networks and Supernetworks: Evolutionarily Conserved Gene Interactions ...................................... 371 Alexander Noveen, Volker Hartenstein, Cheng-Ming Chuong Introduction ......................................................................................... 371 Genes .................................................................................................... 372 Gene Networks .................................................................................... 378 Gene Supernetworks ........................................................................... 381 Summary and Conclusion .................................................................. 386 Part VI: Approaches 20. Current Methods in the Study of Avian Skin Appendages ................. 395 Ting-Xin Jiang, N. Susan Stott, Randall B. Widelitz and Cheng-Ming Chuong Introduction ......................................................................................... 395 Detecting Molecular Expression ......................................................... 395 Skin Explant Cultures .......................................................................... 399 Manipulation of Chicken Embryo ...................................................... 400 Monitoring Cell Behavior ................................................................... 401 Perturbing Functions with Retroviral Mediated Gene Transduction ............................... 403 In Search of Candidate Genes ............................................................. 404 Perspectives .......................................................................................... 406

21. Molecular Biology of Anhidrotic Ectodermal Dysplasia .................... 409 Juha Kere Introduction......................................................................................... 409 The Phenotype of Anhidrotic Ectodermal Dysplasia and Carrier Manifestations ............................................................. 409 Phenotypic Similarity of the Tabby Mouse and Clues for Site of Action ............................................................ 411 Genetic and Physical Mapping of EDA and TA ................................. 411 Cloning of the EDA Gene and Structure of a Predicted Protein ...... 412 EDA Expression and Function ........................................................... 414 22. Systematic Approach to Evaluation of Mouse Mutations with Cutaneous Appendage Defects ..................................................... 421 John P. Sundberg, Xavier Montagutelli and Dawnalyn Boggess Introduction......................................................................................... 421 Systematic Evaluation of New Mouse Mutations .............................. 421 Resources and Repositories ................................................................. 431 Color Insert ...................................................................................................... 437 Index ................................................................................................................ 441

EDITOR Cheng-Ming Chuong Department of Pathology University of Southern California Medical School Los Angeles, California, U.S.A. Chapters 1, 5, 13, 18, 19, 20

CONTRIBUTORS Kathryn D. Anderson Department of Surgery and the Developmental Biology Program Childrens Hospital Los Angeles Research Institute and the Center for Craniofacial Molecular Biology University of Southern California School of Medicine and Dentistry Los Angeles, California, U.S.A. Chapter 12 Dawnalyn Boggess The Jackson Laboratory Bar Harbor, Maine, U.S.A. Chapter 22 Ding Bu Department of Surgery and the Developmental Biology Program Childrens Hospital Los Angeles Research Institute and the Center for Craniofacial Molecular Biology University of Southern California School of Medicine and Dentistry Los Angeles, California, U.S.A. Chapter 12 Yi-Ping Chen Howard Hughes Medical Institute Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts, U.S.A. Chapter 14

Robert deLemos Department of Pediatrics University of Southern California School of Medicine Womens Hospital Los Angeles, California, U.S.A. Chapter 11 Danielle Dhouailly Biologie de la Différenciation Epithéliale Etude de la Différenciation et de l’Adhérence Cellulaires Institut Albert Bonniot Université Joseph Fourier Grenoble, France Chapter 4 Stephanie Dunn Department of Animal Science The University of Adelaide, Waite Campus Glen Osmond, South Australia Chapter 16 Elisabeth Dupin Institut d’Embryologie Cellulaire et Moleculaire du CNRS et du College de France Cedex, France Chapter 8

Guillermo Flores-Delgado Department of Surgery and the Developmental Biology Program Childrens Hospital Los Angeles Research Institute and the Center for Craniofacial Molecular Biology University of Southern California School of Medicine and Dentistry Los Angeles, California, U.S.A. Chapter 12 Margaret Hardy Fallding Department of Biomedical Sciences Ontario Veterinary College University of Guelph Guelph, Ontario, Canada Chapter 3 Volker Hartenstein Department of Molecular, Cellular and Developmental Biology University of California Los Angeles, California, U.S.A. Chapter 19 Ting-Xin Jiang Department of Pathology University of Southern California Medical School Los Angeles, California, U.S.A. Chapter 20 Han-Sung Jung Department of Anatomy and Developmental Biology University College London, London, United Kingdom Chapter 18 Benoit Kanzler Biologie de la Différenciation Epithéliale Etude de la Différenciation et de l’Adhérence Cellulaires Institut Albert Bonniot Université Joseph Fourier Grenoble, France Chapter 4

Juha Kere Department of Medical Genetics Haartman Institute University of Helsinki Helsinki, Finland Chapter 21 Shioko Kimura Laboratory of Metabolism National Cancer Institute Bethesda, Maryland, U.S.A. Chapter 11 Ronit Lahav Institut d’Embryologie Cellulaire et Moleculaire du CNRS et du College de France Cedex, France Chapter 8 Laure Lecoin Institut d’Embryologie Cellulaire et Moleculaire du CNRS et du College de France Cedex, France Chapter 8 Nicole Le Douarin Institut d’Embryologie Cellulaire et Moleculaire du CNRS et du College de France Cedex, France Chapter 8 Richard Maas Howard Hughes Medical Institute Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts, U.S.A. Chapter 14 Parviz Minoo Department of Pediatrics University of Southern California School of Medicine Womens Hospital Los Angeles, California, U.S.A. Chapter 11

Xavier Montagutelli The Institut Pasteur Paris, France Chapter 22 Sven Müller-Röver Department of Dermatology Charitié, Humboldt-Universität zu Berlin Berlin, Germany Chapter 15 Stuart A. Newman Department of Cell Biology and Anatomy New York Medical College Valhalla, New York, U.S.A. Chapter 17 Alexander Noveen Department of Molecular, Cellular and Developmental Biology University of California Los Angeles, California, U.S.A. Chapter 19 Glenn H. Nuckolls National Institure of Arthritis, Musculoskeletal and Skin Diseases National Institutes of Health Bethesda, Maryland, U.S.A. Chapter 2

Ralf Paus Department of Dermatology Charitié, Humboldt-Universität zu Berlin Berlin, Germany Chapters 6, 15 Michael Philpott Department of Anatomy St. Bartholomew’s and the Royal London School of Medicine and Dentistry Queen Mary Westfield College London, England, U.K. Chapter 6 Johanna Pispa Developmental Biology Program Institute of Biotechnology University of Helsinki Helsinki, Finland Chapter 9 Barry Powell Department of Animal Science The University of Adelaide, Waite Campus Glen Osmond, South Australia Chapter 16

Richard E. Olver The Department of Child Health Ninewells Hospital & Medical School University of Dundee Dundee, Scotland Chapter 12

Fabrice Prin Biologie de la Différenciation Epithéliale Etude de la Différenciation et de l’Adhérence Cellulaires Institut Albert Bonniot Université Joseph Fourier Grenoble, France Chapter 4

S. Parimoo Skin Biology Research Center Johnson & Johnson Skillman, New Jersey Chapter 7

Steve Prouty Skin Biology Research Center Johnson & Johnson Skillman, New Jersey, U.S.A. Chapter 7

Drucilla J. Roberts Department of Pathology Massachusetts General Hospital and Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts, U.S.A. Chapter 10 George E. Rogers Department of Animal Science The University of Adelaide, Waite Campus Glen Osmond, South Australia Chapter 16 Lillian Shum National Institure of Arthritis, Musculoskeletal and Skin Diseases National Institutes of Health Bethesda, Maryland, US.A. Chapter 2 Harold C. Slavkin National Institute of Dental Research and National Institute of Arthritis, Musculoskeletal and Skin Diseases National Institutes of Health Bethesda, Maryland U.S.A. Chapter 2 Kurt Stenn Skin Biology Research Center Johnson & Johnson Skillman, New Jersey, U.S.A. Chapter 7 N. Susan Stott Department of Pathology University of Southern California Medical School Los Angeles, California, U.S.A. Chapter 20

John P. Sundberg The Jackson Laboratory Bar Harbor, Maine, U.S.A. Chapter 22 Irma Thesleff Developmental Biology Program Institute of Biotechnology University of Helsinki Helsinki, Finland Chapter 9 Jean P. Viallet Biologie de la Différenciation Epithéliale Etude de la Différenciation et de l’Adhérence Cellulaires Institut Albert Bonniot Université Joseph Fourier Grenoble, France Chapter 4 David Warburton Department of Surgery and the Developmental Biology Program Childrens Hospital Los Angeles Research Institute and the Center for Craniofacial Molecular Biology University of Southern California School of Medicine and Dentistry Los Angeles, California, U.S.A. Chapter 12 Randall B. Widelitz Department of Pathology University of Southern California Medical School Los Angeles, California, U.S.A. Chapters 5, 13, 20

PREFACE

D

r. Landes first approached me to write a book on feather pattern formation. Although it is a topic dear to my heart, after some thought I suggested expanding the topic to cover the theme of epithelial appendages, with reasons stated in my introductory chapter. I then invited experts in related fields to write chapters with the theme of “epithelial appendages” in mind. The response was great. Many feel enthusiastically that it is timely to do so. Indeed, to date, there has not been an effort to organize these topics by researchers examining different epithelial appendages. These fields may appear diverse and yet they do share a common ground: They are all derivatives of epithelia. This book should break the superficial barriers between these fields and bridge the gaps. It is hoped that this book will catalyze the molecular understanding of the fundamental mechanisms of epithelial appendage morphogenesis and help a newly integrated field to emerge. Chapters in this book contain reviews covering recent experimental findings and current thinking on epithelial appendage morphogenesis. Many chapters contain summary diagrams of the formative processes of respective organs, with cellular and molecular explanations. Many chapters also contain useful information on resources. The book is intended for the following constituents: 1. Research scientists directly involved in these areas may find the book informative and helpful to their own research. In addition, the book can provide different and broader perspectives that may give investigators a fresh view of their own research. 2. Graduate students, using it as supplemental reading for developmental biology or other related courses, may find it a stimulating and germinating ground for new ideas. I hope that it will be one, just as Wessel’s book was for me (please see below). 3. Other scientists may find their horizons expanded by giving this book their attention. For instance: • Biomedical research scientists and clinicians may be inspired by the dynamic processes behind tissue and organ formation; • More traditional histologists might see how exciting the study of the molecular mechanisms of tissue design can be; • More traditional molecular biologists might see how rich the study of tissue design and tissue construction can be. 4. For all nature lovers, I hope it is an enjoyable book to glance through and appreciate. Just like a coffee table encyclopedia that shows the beauty and rich diversity of the animal kingdom, this book reveals the unifying force flowing behind tissue diversity. Like music, the morphogenetic process then elaborates by superimposing variations on a common theme. My personal interest in developmental biology stems from my school years. My favorite subjects were embryology and histology. Having an interest in art, I was fascinated by the beautiful and robust changes taking place during embryogenesis and histogenesis. I wondered how Nature created such splendors. Dr. HwaiSan Lin, my histology professor, tried to explain subjects using the best possible mechanistic view. Still, it was obvious that much remained to be learned. In search for answers, I became fascinated by Dr. Wessel’s book “Tissue Interactions and

Development” (Benjamin/Cummings, Menlo Park, CA. 1977) which discusses the classical experiments of epithelial-mesenchymal interactions, including feather development. To satisfy my burgeoning curiosity, I decided to pursue a Ph.D. In 1978, I went to study cell adhesion in embryogenesis with Dr. G. M. Edelman, a Nobel Laureate, at the Rockefeller University in New York. One lesson I learned from his brilliant mind is to look at things from a global view; then simplicity and fundamentals will reveal themselves. During my later years at Rockefeller, I was examining the immunofluorescence of NCAM and NgCAM in developing chicken spinal cord. Suddenly, I saw NCAM light up dermal condensations in the developing skin like stars glowing in the dark night sky. The secondary inductions I read about in Wessel’s book have pointed the way. With Dr. Edelman’s encouragement and support, I obtained Sengel’s “Morphogenesis of Skin” (Cambridge University Press, 1976); I was very inspired by his style, in both thought and approach. I decided to follow the lead of the feather into the spectacular world of embryonic development. In 1987 I came to the University of Southern California and started my own laboratory. In my decade here, I have been pleased to be joined by many colleagues and students in my endeavor. Using the feather as a model, we have surely had a lot of fun figuring out how Nature works in morphogenesis. With the advent of molecular biology, the field has expanded and is ready for a new integrated perspective. It is therefore gratifying that I have the opportunity to assemble this book, reflecting with a unifying theme an intellectual heritage gained from Wessel’s and Sengel’s books, and using the lessons I learned from my Ph.D. mentor, Dr. Edelman. With this background, I would like to thank all contributors for their participation in this project, and for the enormous effort and time they put in. I would like to thank reviewers, and the many colleagues in related fields with whom I have interacted and bothered one way or another. Special thanks go to Dr. Landes and his staff for making this project possible. For me to be in a position to do this job, I am grateful to my laboratory members, long or short term, for their various contributions; particularly Dr. Randall B. Widelitz and Dr. Ting-Xin Jiang, for their dedication and good work, Dr. Alex Noveen for many exciting discussions, and Ms Sheila Delshad for help in editing the manuscripts. From the University of Southern California, I also appreciate the support and encouragement of many colleagues, particularly the Pathology Department chairman Dr. Clive Taylor, vice chairman Dr. Pradip Roy-Burman, and previous Dermatology chairman, Dr. Thomas Rea. For my family, I am grateful to my parents for providing and cultivating an environment that shaped my appreciation of science and nature. Finally, I owe thanks to my wife, Dr. Violet Shen, for her constant understanding, patience and unconditional support. Without these, this book would not be possible. Cheng-Ming Chuong, M.D., Ph.D. Sept. 15, 1997 Department of Pathology University of Southern California

Part I

Overview

CHAPTER 1

Morphogenesis of Epithelial Appendages: Variations on Top of a Common Theme and Implications in Regeneration Cheng-Ming Chuong “To see a world in a grain of sand And a heaven in a wild flower” - William Blake -

The Epithelial Appendage Paradigm

E

pithelial appendages are derivatives of the epithelial sheet. They add structural and functional complexities to the otherwise flat epithelia (Fig. 1.1). Epithelial appendages can be categorized as evaginated or invaginated appendages. Evaginated epithelial appendages like feathers and intestinal villi are appendages that protrude out of the epithelial surface. Invaginated epithelial appendages like mammary glands, lung and bile duct epithelia are appendages that invaginate into the mesenchyme. These epithelial appendages can also be categorized by location. On the body surface are hairs, feathers, scales, claws, beaks, horns, sebaceous glands, sweat glands, mammary glands, etc. In the oral cavity, there are teeth and salivary glands. Inside the body, the gut mucosa is covered with different derivatives that line the stomach, small intestine, colon, etc. During embryonic development, the derivatives of the endodermal tube form many other important organs including the lung, pancreas, liver, etc. Among epithelial appendages, the diversification of skin appendages (or cutaneous appendages) and teeth are most obvious since they form the direct interface between the organism and the environment.2 Their function has gone beyond the original barrier and protective roles. The evolution of different skin appendages has enabled the vertebrate animals to adapt into different niches.1 They have become central to locomotion (feathers for flying, snake scales for crawling), insulation (hairs and down feathers), communication (emotional display, sexual attraction), competition (claws, teeth, beaks), defense (quills, turtle shells), sensation (vibrissae, bristles), etc. In fact, the presence of some appendages has become part of the definition of specific vertebrate classes: i.e., hairs and feathers are key characteristics of mammals and birds, respectively. All of these different three dimensional epithelial appendages are topological alterations of the two dimensional epithelia. What are the driving forces behind these morphogenetic Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

4

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 1.1. A schematic drawing of different epithelial appendages. A variety of epithelial appendages are shown. Some evaginate and some invaginate. Some derive from the ectoderm and some from the endoderm. They are all products of epithelial-mesenchymal interactions based on the same theme.

Morphogenesis of Epithelial Appendages

5

processes? What do these apparently different organs share in common? Why put them under the theme of epithelial appendages? First let us have an overview of the previous studies.

Studies on Epithelial Appendage Morphogenesis Early embryological studies showed that, despite the differences in final products and germ layer origins, epithelial appendages share similar developmental processes. All are the results of epithelial-mesenchymal tissue interactions during embryonic organogenesis. In most cases, mesenchymal tissues acquire specificity information and epithelial tissues elaborate and differentiate to form the appendages per se. The epithelium may be of ectodermal or endodermal origin. The mesenchyme can come from either the somite or neural crest. They share fundamental mechanisms including epithelial folding, cell migration, invagination, differential cell proliferation, branching, etc. The epithelial and mesenchymal components from different epithelial appendages can even be interchangeable, if tested at early stages. Where, then, do the specificities of epithelial appendages come from?

Classical Studies on Morphological Variety and Specificity Investigators have searched for the origin of epithelial appendage variation for a long time. The question was addressed elegantly approximately 2-3 decades ago (summarized in ref. 3). Epithelial appendages are distinct at several different levels: among different classes (e.g., chicken feather and reptile scale); species (chicken and duck feather); locations (hair and teeth); and types (flight and down feather). This was addressed by recombining epithelium and mesenchyme from different sources and allowing them to grow (Fig. 1.2).4,5 In most cases, the mesenchyme determines the phenotype (please see chapter 4 for further discussion). For example, chicken chorioallantoic membrane can be induced to form feathers and scales.6 The message can also go across species. However, epithelia can only respond within their capability, so that recombination of chicken feather mesenchyme and mouse epithelium led to the formation of hairs arranged in a feather pattern. Several works suggest that the specificity of epithelial appendages can be altered by chemicals such as retinoic acid when added at an appropriate time. This is seen in the hairgland7 and scale-feather8 transformation experiments. Based on these results, a two stage hypothesis was proposed: 1. The location of the skin appendage is determined; then the identity of the appendage is determined. In the placode stage, the appendage field is defined but remains flexible as to which type of appendage to make. 2. The morphological specificity becomes determined and irreversible. Recombination experiments using epithelia prior to determination result in appendages with the morphology specified by the mesenchyme. Recombinations using epithelia that have developed beyond the determination stage form appendages with the already determined epithelial phenotypes. This helps to explain why, when leg mesenchyme was recombined with wing epithelium, some feather buds still formed.9 Although these experiments enhance our understanding, the molecular nature of these “messages” remained elusive because too few molecules had been identified at that time.

Common Themes Revealed by Molecular Studies In the past 10 years, there has been remarkable progress in developmental biology. This progress was acknowledged by awarding the 1995 Nobel Prize in Physiology and Medicine to developmental biologists, Drs. Lewis, Nusslein-Volhard and Weischaus. Using Drosophila as a model, they identified many genes important for patterning and morphogenesis.10,11

6

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 1.2. Epithelial appendages can regrow after epithelia and mesenchyme are separated and recombined. This shows the process that is common to all epithelial appendages. First a series of interactions takes place between the epithelia (E) and mesenchyme (M). In the early stages of morphogenesis, phenotypes are still flexible. Gradually, they become irreversibly committed (as represented by the increasing size of arrows). Cross recombination between two types of epithelial appendages (E1, E2) shows that the mesenchyme (M1, M2) has more control over the phenotypes of epithelial appendages, but epithelia can only respond within their competence at the time that stimuli are given. See color insert.

Morphogenesis of Epithelial Appendages

7

Data accumulated since then suggest that most molecules used in constructing vertebrate embryos and epithelial appendages, are homologous to those used in Drosophila morphogenesis. One category of candidate molecules is the Homeobox genes.10,12 Homeobox genes are originally identified from Drosophila homeotic mutants and are involved in determining the appendage phenotypes.13 They contain a homeobox domain that binds DNA. Hox, Msx, and Dlx are all used frequently in epithelial appendage morphogenesis. We found that the expression of Hox genes in the developing skin are body position specific.14 Since the types of epithelial appendages are also body position specific, we proposed the skin Hox code hypothesis, suggesting that skin appendage phenotypes are determined by characteristic Hox expression patterns (reviewed in ref. 14 and chapter 4). Retinoic acid alters morphological specificity probably by perturbing skin Hox codes. Similarly, different patterns of Hox gene expression are found along different regions of the gut (chapter 10),16 suggesting a role in determining the epithelial appendage phenotypes of the gut mucosa. Many signaling molecules appear repetitively during the morphogenesis of various appendages (this book). These include Shh, FGFs, BMPs, etc. Similarly, many adhesion molecules also share in carrying out the morphogenetic processes. These include NCAM, integrins, cadherins, fibronectin, collagen, etc.17 Other than the master switch gene and the final differentiation products, there may not be many organ specific genes. However, the shared genes may play different functions in different scenarios.18,19 Many of the signaling molecules are secretory molecules or transcription factors. Among them, some may act early in the molecular cascade, before the morphological specificity is determined. For example, LEF-1, a nuclear factor, is expressed in both hair and tooth development. Interestingly, transgenic mice with over-expression of LEF-1 led to the ectopic formation of hairs in the gum regions.20 Some molecules may be involved in laying down specific axial orientations. Wnt7a/Lmx and engrailed are found to be involved in dorso- and ventralization of limb bud mesenchyme,21,22 and therefore may be involved in phenotypic determination of dorsal and ventral skin phenotypes (hair vs. foot pad). Some genes may be used repetitively in different stages for different morphogenetic functions. Ectopic expression of these molecules may perturb their physiological functions and lead to unexpected results. Recently, suppression of the BMP pathway in the limb bud by injecting RCAS carrying dominant negative BMP receptors led to small feather buds growing out from the distal end of scutate scales.23 Thus modulation of these molecules may influence the spatial and temporal processes of cell adhesion, proliferation and differentiation, which then results in differences in the final products. In summary, so far studies have shown that the formation of all epithelial appendages share the following stages: 1. Induction stage. Out of the homogeneous epithelial sheet, a domain is defined and cells within the domain become destined to form an appendage. 2. Morphogenesis stage. Together with mesenchymal input, the epithelial sheet in this domain will invaginate or evaginate, coupled with differential growth, branching formation, and other morphogenetic movements to mold a group of cells into a particular shape. 3. Differentiation stage. The organ anlagen now grow in size and differentiate into specific gene products. These changes occur with coordinated growth control. Finally, the epithelial appendages, with unique structures and functions, are put to use by the organism.

Molecular Basis of Epithelial Appendage Morphogenesis

8

What Is in this Book? The Goal of this Book With the above background, the goals of this book are to: • Develop the theme of “epithelial appendages” to include both ectodermal and endodermal appendages. • Compare and contrast the common themes and variations of the morphogenetic processes and epithelial-mesenchymal interactions in different kinds of epithelial appendages. • Achieve an understanding of the molecular mechanisms that drive and modulate the morphogenetic processes. • Catalyze the new wave of research on the molecular basis of epithelial appendage formation.

The Content of this Book The book is divided into six parts. Part I Part I is an overview. In this introductory chapter, I review the past and present status of this field, using materials mainly from skin appendages. On a broader level, the concept of skin appendages is expanded to cover endodermal appendages, and the theme of epithelial appendages is developed. This unifying theme is supported by the recent progress in human diseases, including “ectodermal dysplasia”, in which pathogenesis of several epithelial appendages including hair, sweat glands, and nails are hit simultaneously by a mutation in one gene. Using this as a pivotal point, Dr. Slavkin et al present a global view of the current status and future of the field of epithelial appendages. In a recent meeting keynote speech (Animal Models as Biomedical Tools: Mouse Skin and Hair Mutations, Bar Harbor, Sept. 1996), Dr. Hardy reflected on the excitement of her distinguished hair research career. She recalled anecdotes of hair research in its early years. We put the speech here not as a historic record, but as an inspiration. Part II and Part III Parts II and III are aimed to cover different types of epithelial appendages: with four chapters dealing with the appendages on the body surface and four chapters on teeth and endodermal appendages. Chapter 4, by Dhouailly et al, covers three decades of important research and insightful views on the morphological specificity of cutaneous appendages. Roberts (chapter 10) and Minoo et al (chapter 11) echo the similarities and variations of the endoderm side. Regional specificities within the body are as rich as those on the body surface. The feather chapter by Chuong and Widelitz (chapter 5) analyzes the process in more general architectural terms, presenting criteria for candidate molecules rather than focusing on specific molecules, with the intention that some of these principles may be modified for use in the design of other organs. The chapters on hairs, teeth and lung epithelia, by Philpott and Paus, Theslef and Pispa, and Warburton et al respectively (chapters 6, 9 and 12), are the best examples of how different epithelial appendages share the same molecules in different organs. One characteristic of epithelial appendages is that they go through renewal cycles. Stenn et al in chapter 7 present an original discussion of the concept of biological rhythms, from cell cycle to diurnal cycle to seasonal cycle. One of the most dramatic features of the animal integument is the pigmentation pattern. The pattern can be distributed as stripes or dots along the whole body such as in the

Morphogenesis of Epithelial Appendages

9

zebra, or along an epithelial appendage such as in a feather. These patterns are set by the melanocytes. Chapter 8 by Dr. Le Douarin’s discusses the new progress on growth control of melanocytes, one of the derivatives of neural crest cells. Part IV Part IV focuses on molecular mechanisms rather than organs. The cellular process of induction and mesenchymal condensation is discussed in chapter 13 by Widelitz and Chuong. The process involves a series of interactions among signaling molecules and adhesion molecules. Signaling loops in tooth morphogenesis, revealed by combining genetic studies with experimental embryology, are discussed by Chen and Maas in chapter 14. Adhesion molecules, demonstrated by dynamic changes during hair follicle morphogenesis, are presented in the comprehensive chapter 15 by Müller-Röver and Paus. These processes conform with Dr. Edelman’s concept of topobiology.24 To finish the construction of an epithelial appendage, late differentiation events have to be coupled to morphogenesis to consolidate the morphology and fill the anlage with specialized products. Chapter 16 by Rogers et al uses keratinocyte differentiation as an example to illustrate this process. Part V In this age of molecular biology, searching for molecular bases is important. However, too much emphasis on molecular cues in patterning can lead to the expectation of a certain rigid molecular specificity that may not exist. Some morphogenetic processes may result from the interactions and equilibrium of the physical chemical properties conferred to the cells. Of course cells are made up of molecules. However, morphogenesis occurs at the level of cells and tissues, rather than molecules. By focusing on the key principles, Newman’s chapter 17 is a stream of fresh ideas that deserves more weight in the thinking of developmental biologists. The periodic patterning of integument appendages has been a favorite subject for theoretical biologists; however, no molecular basis has been established. Incorporating recent experimental findings, Jung and Chuong present in chapter 18 a feather patterning model based on the Turing model and suggest some compelling molecular candidates to support the model. During the history of evolution, new types of epithelial appendages emerge. There is constant pressure to produce “better tools” for survival. Once invented and proven useful, the species with the new appendage has a “patent”, and is awarded a period of dominance until the environment changes or better appendages are evolved by other species. The sharing of molecules in morphogenetic processes provides a physical basis on which duplication and mutation of gene networks can be built. On this background, a hierarchy of levels of gene network organization is presented by Noveen et al in chapter 19. The fact that the number of epithelial appendages is so great, and alterations less likely to be lethal, may account for epithelial appendages being more “evolvable”.25 Part VI Methodology and resources are basic to epithelial appendage research. Here Jiang et al present in chapter 20 a summary of useful approaches currently used to study chicken embryos, particularly epithelial appendages. Since changes in the skin are easily recognized, more and more scientists originally studying molecules not known to be related to skin appendages have observed interesting phenotypes in the skin of transgenic mice. Sundberg et al’s chapter 22 provides a very useful primer on further analyses of these transgenic mice.

10

Molecular Basis of Epithelial Appendage Morphogenesis

As stated above, with the progress in molecular genetics and the Human Genome Project, one of the major advances in the next phase of epithelial appendage research will come from the identification of many genes involved in ectodermal dysplasia and other related human or mouse genetic disorders. Kere’s chapter 21 describing his identification and initial characterization of EDA sets a pioneering example for us to follow.

Organogenesis, Histogenesis, and New Growth Organogenesis is the study of the process of organ formation. Other than during development, it also happens during the cycling of epithelial appendages and regeneration. Hairs and feathers go through molting cycles. In each cycle, dermal papilla cells induce epithelial stem cells to reform a new appendage. In teeth, humans only have one cycle, but other animals can cycle repetitively. Following injury, healing with re-epithelialization and connective tissue repair fulfills the minimal basic requirement. It has long been hoped that someday damaged tissue can be replaced with skin that has functional appendages. Cloning of a whole animal is one possibility but has its problems.26 An alternative is to have the ability to initiate or modulate organogenesis from primary cells or tissue fragments obtained from oneself. This will require knowledge in three fundamental processes: 1. identification of stem cells and the molecular nature of their competence; 2. identification of regulators of morphogenesis, or how the three dimensional arrangement of the cells and tissue are assembled; and 3. identification of regulators that control the proper differentiation of these structures to assume specialized functions. To achieve this, two problems must first be solved. One is to identify and expand the pool of stem cells. The other is to instruct these cells to undergo proper morphogenetic processes. The search for epithelial stem cells and hair stem cells has used different approaches based on the properties of stem cells. These include slow cell cycling in vivo,27 an ability to become clonogenic in vitro,28 and expression patterns of adhesion molecules.29 Because of these differences, it is sometimes debatable which is the real stem cell.30 We could choose to use a developmental biology perspective and use the term “epithelioblast” or epithelia precursor, which is less stringent but probably reflects a more physiological condition (Fig. 1.3). By definition, stem cells should have two properties: self renewal and pluripotentiality. However, these abilities do not come and go abruptly; rather, they are gained or lost progressively during development, and become irreversible only after passing an unknown determination stage. Cells in these stages can be termed “epithelioblast” or “keratinoblast”, paraphrasing terms used in myoblast, neuroblast, erythroblast, etc. Current research should aim to define different stages of epithelioblasts. These can be defined based on their pluripotentiality, molecular markers, and the properties that come with those molecules (for an initial example see ref. 31). If we can learn how signals induce epithelioblasts from one stage to the other, or de-differentiate, to the level of our understanding of how cytokines influence blood stem cell differentiation,32 we may have a better handle to manage disease conditions involving epithelia and epithelial appendages. We may also be in a better position to identify and isolate stem cells (or the earliest epithelioblast) from adult tissues, or to make later stage (partially differentiated) epithelioblasts de-differentiate and become more pluripotential. With the aid of gene therapy and appropriate tissue engineering, we should be able to reproduce some morphogenetic processes and to regenerate organs that are at least partially functional. Epithelial appendages are made up of more than one histology type. From studies on the morphogenesis of epithelial appendages, we realize how little we know about what determines basic histology types such as stratified epithelia, squamous epithelia, cuboidal epithelia, or transitional epithelia. A change from simple epithelia to multiple layered epithelia

Morphogenesis of Epithelial Appendages

11

Fig. 1.3. A proposed scheme of different epithelioblast stages. This shows progression of epithelial cell (E) differentiation. A and B represent different types of epithelial appendages. The numbers represent different developmental stages. E1 is the ultimate epithelioblast or stem cell. E6 is the irreversibly differentiated keratinocyte. From E2 to E5 are different stages of epithelioblast. Although E2 and E3 are not stem cells, they are multi-potential epithelioblasts and can de-differentiate back to stem cell status if appropriate signals and environments are provided. The plasticity gradually diminishes, as represented by the decreasing size of the reverse arrows. In appendages A and B, A1-A3 represent inductive, morphogenetic and differentiation stages respectively, as described in the text. The same holds for B1-B3. The reversibility decreases gradually as cells become more differentiated. The difference in competence observed in cells from the hair bulge, matrix and outer root sheath30 may represent a mixture of cells with defferent levels of competence. x, y, and z represent signaling molecules at key branching points, whose appearance or disappearance are important for directing the fate of epithelial cells. Examples are Shh and FGF. Different epithelioblast stages (designated by numbers) probably have different molecular markers which need to be defined. Some may be expressed only in E1, while some may be expressed transiently from E2 to E3.

would mean the absence of contact inhibition in the monolayer and involve a 90o turn of the mitotic spindle axes. How are these events regulated in normal histogenesis? Are all these also under mesenchymal control? In a practical sense, normal growth of an epithelial appendage represents regulated new growth, while cancer is new growth that goes out of control. Epithelial-mesenchymal interactions also play an important role in breast and prostate carcinoma.33 Metaplasia is an early sign of cancerous changes, but formation of new histological phenotypes is a normal process in development. Increases in epithelial mobility can be a prelude to metastasis, but cell migration is also a normal process in morphogenesis.

12

Molecular Basis of Epithelial Appendage Morphogenesis

Therefore by understanding the normal function and regulation of cancer related genes in embryo development, we may learn some useful lessons and new perspectives on the management of cancer. In this context, the study of epithelial appendages has a broader significance: in development, regeneration and cancer. By searching for the similarities and specificities among these different appendage organs, we appreciate a more unifying view on development and evolution. We also come to a more fundamental understanding of the molecular mechanisms underlying morphogenesis, organogenesis, and pathogenesis. Since epithelial appendages such as hair follicles go through constant renewal and induction during adult life, they must have a higher percentage of epithelioblasts and higher competence for regenerative processes. Although tiny, a hair follicle is a rich source of information and power. This reminds me of the Chinese legend “A Journey to the West”, which was written about one thousand years ago. A mythological monkey has to escort his master from China to India to bring back Buddhist books, despite all the physical difficulties of that time. One unique trait this monkey has is that he can actually pluck off some of his body hairs, and turn them into clones of himself with one magical blow. We need to know the molecular basis of this magic stroke.

Acknowledgment This work is supported by grants from NIH and NSF. I thank Dr. Randall B. Widelitz and Mr. Edward B. Chuong for help in computer drawing of Figs. 1.2 and 1.3, respectively.

References 1. Burton M. The New Larousse Encyclopedia of Animal Life. New York: Bonanza Books, 1967. 2. Bereiter-Hahn J, Matoltsy AG, Sylvia Richards K. Biology of the Integument. Vol. 2. Berlin: Springer-Verlag, 1984. 3. Sengel P. Morphogensis of Skin. Cambridge: Cambridge University Press, 1976. 4. Saunders JW, Gasseling MT. The origin of pattern and feather germ tract specificity. J Exp Zool 1957; 135:503-528. 5. Dhouailly D. Formation of cutaneous appendages in dermo-epidermal interactions between reptiles, birds and mammals. Roux Arch Dev Biol 1975; 177:323-340. 6. Dhouailly D. Feather-forming capacities of the avian extra-embryonic somatopleure. J Embryol Exp Morphol 1978; 43:279-287. 7. Hardy MH. Glandular metaplasia of hair follicles and other responses to vitamin A excess in cultures of rodent skin. J Embryol Exp Morphol 1968; 19:157-180. 8. Dhouailly D, Hardy MH, Sengel P. Formation of feathers on chick foot scales: a stagedependent morphogenetic response to retinoic acid. J Embryol Exp Morph 1980; 58:63-78. 9. Sengel P, Pautou MP. Experimental conditions in which feather morphogenesis predominates over scale morphogenesis. Nature 1969; 222:693-694. 10. Lewis EB. A gene complex controlling segmentation in Drosophila. Nature 1978; 276:565-570. 11. Nusslein-Volhard C, Wieschaus E. Mutations affecting segment number and polarity in Drosophila. Nature 1980; 287:795-801. 12. Krumlauf R. Hox genes in vertebrate development. Cell 1994; 78:191-201. 13. Shubin N, Tabin C and Carroll S. Fossils, genes and the evolution of animal limbs. Nature 1997; 388:639-648. 14. Chuong CM, Oliver G, Ting SA et al. Gradients of homeoproteins in developing feather buds. Development 1990; 110:1021-1030. 15. Chuong C-M. The making of a feather: Homeoproteins, retinoids and adhesion molecules. BioEssays 1993; 15:513-521.

Morphogenesis of Epithelial Appendages

13

16. Roberts DJ, Johnson RL, Burke AC et al. Sonic hedgehog is an endodermal signal inducing Bmp-4 and Hox genes during induction and regionalization of the chick hindgut. Development 1995; 121:3163-3174. 17. Jiang TX, Chuong CM. Mechanism of skin morphogenesis. I. Analyses with antibodies to adhesion molecules tenascin, N-CAM, and integrin. Dev Biol 1992; 150:82-98. 18. Gehring WJ. The master control gene for morphogenesis and evolution of the eye. Genes Cells 1996; 1:11-15. 19. Piatigorsky J, Wistow G. The recruitment of crystallins: new functions precede gene duplication. Science. 1992; 252: 1078-1079. 20. Zhou P, Byrne C, Jacobs J et al. Lymphoid enhancer factor 1 directs hair follicle patterning and epithelial cell fate. Genes Dev 1995; 9:700-713. 21. Parr BA, McMahon AP. Dorsalizing signal Wnt-7a required for normal polarity of D-V and A-P axes of mouse limb. Nature 1995; 374:350-353. 22. Loomis CA, Harris E, Michaud J et al. The mouse Engrailed-1 gene and ventral limb patterning. Nature 1996; 382:360-363. 23. Zou H, Niswander L. Requirement for BMP signaling in interdigital apoptosis and scale formation. Science 1996; 272:738-741. 24. Edelman GM. Topobiology: An Introduction to Molecular Embryology. New York: Basic Books, 1988. 25. Gerhart J, Kirschner M. Cells, Embryos and Evolution. London: Blackwell Scientific,1997. 26. Wilmut I, Schnieke AE, McWhir J et al. Viable offspring derived from fetal and adult mammalian cells. Nature 1997; 385:810-813. 27. Cotsarelis G, Sun TT, Lavker RM. Label-retaining cells reside in the bulge area of pilosebaceous unit: implications for follicular stem cells, hair cycle, and skin carcinogenesis. Cell 1990; 61:1329-1337. 28. Rochat A, Kobayashi K, Barrandon Y. Location of stem cells of human hair follicles by clonal analysis. Cell 1994; 76:1063-1073. 29. Jones PH, Harper S, Watt FM. Stem cell patterning and fate in human epidermis. Cell 1995; 80:83-93. 30. Reynolds AJ, Jahoda CA. Hair follicle stem cells: characteristics and possible significance. Skin Pharmacol 1994; 7:16-19. 31. Chuong CM, Widelitz RB, Ting-Berreth S et al. Early events during avian skin appendage regeneration: Dependence on epithelial-mesenchymal interaction and order of molecular reappearance. J Invest Dermatol 1996; 107:639-646. 32. Spangrude GJ, Heimfeld S, Weissman IL. Purification and characterization of mouse hematopoietic stem cells. Science 1989; 244:1030. 33. Hayward SW, Cunha GR, Dahiya R. Normal development and carcinogenesis of the prostate. A unifying hypothesis. Ann NY Acad Sci 1996; 784:50-62.

CHAPTER 2

Ectodermal Dysplasia: A Synthesis Between Evolutionary, Developmental, and Molecular Biology and Human Clinical Genetics Harold C. Slavkin, Lillian Shum and Glen H. Nuckolls

I

n 1875 Charles Darwin reported a fascinating set of phenotypes associated with the “toothless men of sind”, members of a Hindu kindred which resides in the vicinity of Hyderabad in India.1 Darwin described a family “in which ten men, in the course of four generations, were furnished, in both jaws taken together, with only four small and weak incisor teeth and with eight posterior molars. The men thus affected have very little hair on their body, and become bald early in life. They also suffer much during hot weather from excessive dryness of the skin. It is remarkable that no instance has occurred of a daughter being thus affected...though the daughters in the above family are never affected, they transmit the tendency to their sons; and no case has occurred of a son transmitting it to his sons.” What Darwin described, prior to the recognition of Mendelian genetics, was the condition now called anhydrotic ectodermal dysplasia (EDA); one of more than 150 distinct forms of ectodermal dysplasia, so-called because the clinical phenotypes appear in tissues derived from the ectoderm. In EDA, affected males show anhydrosis, hypotrichosis and hypodontia. Meanwhile, cellular, developmental, molecular and evolutionary biology blossomed in the 20th century, and have forged an intellectual synthesis between the biological and clinical health sciences. These advances, coupled to numerous technological advances with recombinant DNA, instrumentation and informatics, further accelerated progress towards identifying and characterizing specific human gene mutations associated with disease. In August, 1996, a team of investigators reported the site of the mutated gene for EDA located on the human X chromosome. This unique gene appears to be a transmembrane protein expressed on a number of tissues associated with EDA phenotypes. These and related studies now provide unique scientific opportunities to increase the knowledge base for ectodermallyderived epithelial appendages and to provide the essential information necessary to promote health, reduce disease, and improve both diagnostics and “smarter” therapeutics.

Introduction The ectodermal dysplasias comprise a large and quite heterogeneous group of conditions to which new entities are added each year. In 1984 Newton Freire-Maia and Marta Pinheiro published a comprehensive review that provided an in-depth and then current Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

16

Molecular Basis of Epithelial Appendage Morphogenesis

understanding of both ectodermal dysplasias and the ectodermal dysplasia/malformation syndromes.2 Their published review constitutes an invaluable diagnostic reference work for that time in recent history. Coincidentally, in 1984, several scientific teams published the discovery of homeobox genes in the fruit fly as well as in mammals.3-5 Importantly, a number of discoveries were also published associated with cell and substrate adhesion molecules on the cell surfaces of many cell types,6-8 the paradigm for combinatorial processes in many biological systems,9-10 the discovery of several growth factors and their cognate receptors,11-16 and a series of planning meetings towards the realization of the Human Genome Project in 1988. Now, almost 14 years later, we have experienced a remarkable series of discoveries in the evolutionary, developmental and molecular biological sciences which have profoundly contributed toward understanding the molecular basis for the developmental processes associated with invertebrate and vertebrate embryogenesis, with particular progress towards understanding transcriptional controls for cephalic-caudal and dorsal-ventral developmental gradients, signal transduction process, apoptosis or programmed cell death mechanisms, and a number of molecular features of evolutionary processes. These and other advances are all relevant towards understanding the cluster of diseases and disorders termed ectodermal dysplasias. The descriptions of three germ layers (ectoderm, mesoderm and endoderm) and their fate have evolved through scientific advances in the molecular basis of transcriptional controls, translation and post-translational controls, epigenetic signaling, and a variety of intracellular pathways that connect the extracellular environment with nuclear functions. A number of investigations clearly indicate that many transcription factors (e.g., high-affinity DNA binding proteins, Hox and homeotic proteins), apoptotic regulatory proteins, intracellular and transmembrane receptors, a number of growth factors and cytokines, and a host of unique motifs found within extracellular matrix proteins, are expressed by ectoderm-, mesoderm- and endoderm-derived cell types at various times, positions and stages of development. A number of morphoregulatory molecules are now known to be transiently expressed in each of the three germ layers during embryogenesis. Further, evidence indicates that many of these morphoregulatory molecules are utilized in different combinations and that the diversity of these combinations might provide the specificity for many diverse controls during human embryogenesis. Moreover, the expression of many of these morphoregulatory molecules are highly conserved throughout invertebrate as well as vertebrate evolution—in fruit flies, round worms, the early chordate speciation of 500 million years ago and throughout subsequent evolution. During early vertebrate embryogenesis, such as determination of the anterior-posterior axis within initial neurulation, ectoderm gives rise to the neuroectoderm of the forebrain, midbrain and hindbrain and the ectoderm-derived epithelial tissue that serves to cover the exterior of the forming embryo. Importantly, in those regions within the neuroectoderm of the brain or the epithelial covering of the embryo which are associated with instructive cell-cell interactions (e.g., branchial arch formations, sensory placodes, odontogenic placodes, zone of polarizing activity in limb development, and comparable putative sites for hair, feather and other epidermal appendage morphogenesis), comparable combinations of morphoregulatory proteins are expressed (e.g., Msx-1, Msx-2, Dlx2, Dlx5, Shh, PTC, BMP-2, BMP-4, and FGF-4, FGF-8 and their cognate receptors, integrins, cadherins) (for recent reviews, see refs. 17-20). These discoveries, mostly published since 1984, have been augmented through a strategy to analyze the function of one or several morphoregulatory genes by testing their effects in conditions of under-, over-, or misexpression and null mutation transgenic animal models (e.g., zebrafish, mouse), or using explants in tissue or organ culture. First, investigators

Ectodermal Dysplasia

17

demonstrate the transient expression of the candidate morphoregulatory gene product; for example, a homeotic transcription factor Msx-2 is transiently expressed in rhombomeres within the forming hindbrain, cranial sutures, first branchial arch, tooth organs, heart and limb.21,22 Second, transgenic animal models are created to determine a cause and effect relationship for Msx-2; based upon the patterns of gene expression during embryogenesis, a number of clinical phenotypes can be predicted from either targeted overexpression or disruption of Msx-2.23,24 Possible outcomes could show essentially no apparent clinical phenotype. In the case of no clinical phenotype, scientists often interpret this result from a null mutation to suggest that the specific morphoregulatory gene shares redundant functions with other genes—so called “developmental redundancy”. In other examples, the results from null mutations can be counter-intuitive in that the clinical phenotype is presented in some but not all of the tissues that previously showed transient expression. For example, Msx-1 is expressed in cardiac tissues but does not appear to produce heart defects in null mutation transgenic mice; rather the animals are born with cleft secondary palate and tooth agenesis.25 Another discovery has been the demonstration of transdifferentiation being more common during vertebrate development than previously appreciated. Transdifferentiation describes events when one embryological germ layer or tissue type transforms or transdifferentiates into a different germ layer or tissue type (for recent reviews, see refs. 26-29). Examples are ectoderm giving rise to mesoderm, hindbrain neuroepithelium giving rise to cranial neural crest cells (ectomesenchyme cells and their derived cell lineages), trunk neural tube neuroepithelium giving rise to several phenotypes including melanoblasts, and secondary palatal epithelium transdifferentiating into mesenchymal cells during palatal fusion along the midline of the forming craniofacial complex and kidney development. Human clinical genetics has also provided a number of recent discoveries that change how we consider simple and complex human genetic diseases such as ectodermal dysplasia.30 For example, different mutations in the extracellular domain of the fibroblast growth factor receptor-2 results in five different craniofacial syndromes—Crouzon’s, Apert’s, Jackson-Weiss, Pfeiffer and Beare-Stevenson syndromes. Small but different mutations in the same region of the same gene confer different clinical phenotypes (for reviews, see refs. 31-33). Molecular characterization of inherited human diseases can identify critical residues in proteins that present themselves as sites for DNA binding, protein-protein association, helical turns, phosphorylation, disulfide linkage, proteolytic cleavage, lipid modification, glycosylation and ubiquination. The concerted effort of cell, developmental and molecular biologists and clinical geneticists are building opportunities to bridge the knowledge gap between genotype and phenotype, so as to improve the prevention, detection, diagnosis and treatment of human diseases. In this review, we illustrate ectodermal dysplasias as examples of complex genetic diseases to demonstrate a new synthesis between evolutionary, developmental, and molecular biology and human clinical genetics.

Ectodermal Dysplasias Ectodermal dysplasias (EDs) represent a heterogenous group of genetic disorders that are identified by the absence or defect in two or more of the cardinal signs: skin, hair, nails and teeth. Classification of different forms of EDs reflect over 150 clinically recognized conditions.2,34 The literal definition of “ectodermal dysplasia” describes developmental defects affecting the ectoderm. The ectoderm is one of three germ layers forming the embryo proper. Multiple tissues are derived from or composed of this layer, including skin, skin appendages (hair follicle, nail, sebaceous glands, sweat glands), melanocytes, mammary glands, pituitary gland, teeth, inner ear, optic lens, retina, central nervous system, cranial sensory ganglia

18

Molecular Basis of Epithelial Appendage Morphogenesis

and nerves, skeletal elements of the branchial arches, head mesenchyme, pineal body and adrenal medulla. According to Weech in 1929,35 EDs can be so designated if they include the following features: 1. disturbances that affect tissues of ectodermal origin; 2. disturbances that are developmental; and 3. heredity playing an important causal role. It is only by convention that the clinical definition of ED assumed the four cardinal signs as the diagnostic criteria of defects in hair, nails, teeth and sweat glands. EDs known as Group A conditions involve two or more of the cardinal signs and are classified into 11 clinical subgroups. However, there are numerous other conditions, such as Group B, that involve one of the cardinal signs and defects occurring in another ectodermallyderived tissue. These conditions range from mild to life-threatening. In particular, due to the absence or reduced number of sweat glands, patients can suffer from inability to heat regulate, hyperthermia and complications such as brain damage. Sweating can also be excessive or patchy. These defects in sweating can lead to multiple cutaneous lesions due to dryness, cracking, infections and verrucous eruptions. Primary skin defects can include hyperpigmentation, mosaic pigmentation, keratosis, papillomas, ulcers, bullae, cysts, acrocyanosis, erythroderma, desquamation, atrophy, pterygium, wrinkling, telangiectases or cornification. Nail is characteristically thickened, discolored, brittle, grooved, striated, dystrophic, possessed of celonychia, split or absent. Nail growth is generally slow, with fetid suppuration. Hair defects can range from hypotrichosis, baldness, alopecia, fuzzy, coarse, fine, dry or hypochromic hairs or abnormal hairline affecting body hair, eyebrows and lashes. Teeth defects are also a kaleidoscopic collection of adontia, hypodontia, hypoplasia, supernumerary teeth, irregular placements, natal teeth, delayed eruptions, persistence of deciduous teeth, mis-shaped teeth, discoloration, striations or carious degenerations. In addition to skin, hair, nails and teeth, a wide range of organs and tissues may display dysmorphogenesis such as cleft lip/palate, deficient tears, deficient saliva, deficient mucosal glands of the respiratory track, anomalies in craniofacial morphometry, mental retardation, growth retardation, psychomotor impairment, leukoplakia, anemia, congenital heart disease and immunodeficiencies, and defects in hearing, vision, limbs, mammary glands, urogenital system and skeletal system. The inheritance of EDs suggests that the majority represent either autosomal dominant or recessive genes. Among the numerous and diverse conditions of EDs, only a handful have an identification of a mutated gene mapped to a specific human chromosomal location (Table 2.1). Of the eight known X-linked conditions, five have a chromosomal assignment of a mutated gene. The ED first described by Charles Darwin, is anhydrotic ectodermal dysplasia (EDA; OMIM #305100; ref. 36), an X-linked condition discussed in further detail in the next section and in chapter 21.

Anhydrotic Ectodermal Dysplasia (EDA) EDA, also commonly known as hypohydrotic ectodermal dysplasia (HED), or ChristSiemens-Touraine (CST) syndrome, is an X-linked recessive trait affecting up to 7 in 10,000 live births, the most prevalent form of ED (communicated by the National Foundation of Ectodermal Dysplasia, Mascoutah, Illinois). Over 120 years after the first description of EDA by Charles Darwin in 1875, the gene in which disruptions are associated with the syndrome has been isolated.35 The identification of the gene was based on previous key findings, physical mapping and linkage analyses that suggested that the EDA gene was located on chromosome X between markers q12.2 and q13.1.37-47 Subsequent fine mapping and positional cloning yielded genomic clones which were sequenced and assayed for open reading frames. Primers and probes for the expressed

Ectodermal Dysplasia

19

Table 2.1. Ectodermal dysplasias for which chromosomal location of the gene has been mapped Condition

anhydrotic ectodermal dysplasia; EDA; hypohydrotic ectodermal dysplasia; HED; XHED; Christ-Siemens-Touraine syndrome; CST syndrome partial anodontia; hypodontia tooth agenesis; selective tooth agenesis, familial pachyonychia congenita, Jadassohn-Lewandowsky type; Jadassohn-Lewandowsky syndrome pachyonychia congenita, Jackson-Lawler type; PCHC1

dyskeratosis congenita; ZinsserCole-Engman syndrome incontinentia pigmenti, type I, sporadic type; IP1; BlochSulzberger syndrome incontinentia pigmenti, type II, familial, male-lethal type; IP2 focal dermal hypoplasia; DHOF; FODH; FDH; Goltz syndrome; Goltz-Gorlin syndrome hydrotic ectodermal dysplasia; HED; Clouston syndrome trichorhinophalangeal syndrome, type I; TRPS1 EEC syndrome; ectrodactyly ectodermal dysplasia and cleft lip/palate Ellis-Van Creveld syndrome; chondroectodermal dysplasia; mesoectodermal dysplasia metaphyseal chondrodysplasia, McKusick type; cartilage-hair hypoplasia; CHH Rothmund-Thomson syndrome; RTS; Poikiloderma atrophicans and cataract

MIM Chromosomal Number* Location (Gene Cloned) 305100 Xq12.2-q13.1 (EDA) Kere et al, 1996

106600

4p16.1 (MSX1) Vastardis et al, 1996

Trans- Status§ mission† XR

C

AD

C

167200

17q12-q21 (keratin-16) McLean et al, 1995

AD

C

167210

AD

C

305000

17q12-q21 (keratin-17) McLean et al, 1995 or 12q12-q14 (keratin 6A) Bowden et al, 1995 Xq28

XR

C

308300

Xp11.21

XD

C

308310

Xq28

XD

C

305600

Xp22.31

XD

P

129500

13q

AD

P

190350

8q24.12

AD

P

129900

7q11.2-q21.3

AD

T

225500

4p16

AR

P

250250

9p13

AR

P

268400

8

AR

T

* OMIM, 1996 (ref. 134) † XR, X-linked recessive; XD, X-linked dominant with male lethality; AR, autosomal recessive; AD, autosomal dominant § C, confirmed; P, provisional; T, tentative

20

Molecular Basis of Epithelial Appendage Morphogenesis

open reading frames were used to screen a salivary gland cDNA library to obtain the EDA cDNA. This completely novel cDNA is 854 bp in length encoded by two exons separated by a 200 kb intron. The predicted protein is 135 amino acids in length, belonging to the type II integral membrane proteins, a small class of transmembrane proteins for which very little functional data is known. These proteins contain a signal-anchor domain, so that the polypeptide is inserted into the membrane with the N-terminus cytoplasmic and the C-terminus extracellular. Exon 1 of the gene encodes all but the last three amino acids of the EDA protein. Northern analyses reveal that the 6 kb mRNA of EDA is expressed in a wide variety of fetal and adult tissues including brain, lung, liver, kidney, heart, placenta, skeletal muscle and pancreas. In situ hybridization localized EDA expression in the epidermis, basal cell layer of sebaceous glands, sweat glands, epithelial cells of the upper hair follicle, hair matrix and outer root sheath of the lower hair follicle. EDA expression correlates with those tissue types that are defective in EDA syndromes, suggesting that the EDA protein does play an instructive role in the development of these ectodermally-derived tissues. Sequence analyses of 15 patients afflicted with EDA revealed that six had X:autosomal translocation or deletion, and nine had single base point mutations, including two insertions, two deletions and five substitutions. One recurring mutation was the substitution of leucine to arginine at position 69, suggesting that this amino acid may have a particularly significant function in the protein. In addition, mutation in a neighboring amino acid, a tyrosine to histidine substitution at position 61, was detected in another patient, suggesting that the region encompassing these positions may be an important functional domain. Since gross chromosomal rearrangements, mutations causing frame shifts and protein truncations, and point mutations all give rise to the same phenotype, it is presumed that the mutations are loss of function. The genomic sequence of EDA contains a motif, HK-1,48 positioned upstream of the 5' end of EDA cDNA.36 This regulatory sequence appears to be specific to the promoters of hair follicle genes, such as the keratin intermediate filaments and the keratin associated proteins,48 and this motif was shown to bind to lymphoid enhancer factor 1, Lef1.49 Further support for the genetic interaction of Lef1 and EDA comes from the ED-like phenotype of the Lef1 null mutation in mice (discussed below).

Other Human ED Genes Agenesis of one or more teeth is one of the most common human developmental anomalies, occurring in approximately 20% of the population.50 The inherited condition called familial tooth agenesis (FTA) can be transmitted as dominant, recessive or X-linked. The best characterized form of FTA (OMIM #106600; ref. 51) is autosomal dominant, arising from a point mutation in the Msx-1 gene; although Msx-1 mutations may only account for a small subset of FTA syndromes. Msx-1-FTA is primarily characterized by the absence of the second premolars, maxillary lateral incisors and third molars, with a range of variability in number and location of teeth involved. Since this syndrome often manifests brittle nails, delayed hair growth and delayed teething, it is also classified under EDs.52 The reported mutation causes an arginine to proline substitution in the homeodomain, a highly conserved residue in a structure that interacts with other proteins and with the target DNA molecule. Msx-1 is a homeobox-containing transcription factor that is expressed and functionally active in various embryonic tissues that undergo epithelial-mesenchymal interactions, such as the dental epithelium and mesenchyme, the progress zone of the developing limbs, and nail and hair follicles.53-56 In addition, it is also expressed in a temporally and spatially restrictive manner in the developing rhombomeres57 and their respective neural crest ectomesenchyme, but not in migratory neural crest cells. The function of Msx-1 is not yet clear, although it is thought to play a significant role in regulating the proliferation, differentiation or apoptotic elimination of cells during the morphological development of

Ectodermal Dysplasia

21

the limb and craniofacial regions as well as other key sites of epithelial-mesenchymal interactions in the body. The targeted disruption of the Msx-1 gene in mice generates no defects in heterozygotes; however, the homozygotes exhibit multiple craniofacial defects including complete tooth agenesis, cleft secondary palate, deficiencies in alveolar bone of the mandible and maxilla and abnormalities of the maleus. Tooth initiation occurs in the Msx-1 mutant mice, but the tooth development program is arrested early, and teeth are replaced by bone.25 Since FTA patients exhibit phenotypic features similar to that of the Msx-1 null mutant mice, these patients may have diminished Msx-1 activity. Similar to arrest of tooth development in the mouse, a human patient with the Msx-1-FTA was found to have one second premolar nonerupted and delayed in development and all other second premolars absenct. Since features of the Msx-1 null mice are more severe, some Msx-1 activity may be retained in the FTA patients. The proline for arginine substitution in Msx-1 may create a dominant negative mutant that competitively interferes with the interaction of the wild-type molecule with target DNA sites or with other proteins. Msx-1 has been found to form homodimers and heterodimers with members of the Dlx homeodomain family of proteins, interacting through residues in the homeodomain.58 Further biochemical analysis may clarify the effects of the homeodomain mutation in FTA on the interaction of the Msx-1 molecule with DNA target sites or other homeodomain molecules. Many of the sites of Msx-1 expression overlap or are adjacent to sites of expression of Msx-2. Like Msx-1, mutations in Msx-2 are associated with human disease, since a proline to histidine substitution at position 7 of the homeodomain in Msx-2 causes craniosynostosis, Boston type.59 In FTA, Boston type craniosynostosis, as well as in the Msx-1 null mutation in mice, many of the sites of expression of Msx-1 and 2 appear unaffected by these mutations, which suggests functional redundancy. The overlap of function of these gene products is further suggested by the phenotype of the Msx-1/2 double null mutations, which have defects in almost all organ systems, extensively more severe than the combined phenotypes of the individual null mutations (presented by R. Maas at Keystone Symposium: Bone, Cartilage and Collagen, 1997, Santa Fe, NM). Contrary to these findings, whole mouse embryos in culture treated with a combination of Msx-1 and Msx-2 antisense oligonucleotides exhibited no greater malformations than those treated with either oligonucleotide individually.60 Differences between the results from these two experimental model systems may be due to the window of time of exposure to oligonucleotides, or to limited access of the oligonucleotides to different regions or compartments of the embryo in culture. Two related syndromes are attributed to mutations in keratin subtypes. JadassohnLewandowsky type pachyonychia congenita (OMIM #167200) is associated with substitution in keratin 16 (leucine 132 to proline)61 or keratin 6A (deletion).62 Jackson-Lawler type pachyonychia congenita (OMIM #167210) is due to mutations in keratin 17 (asparagine 92 to serine or aspartate, or tyrosine 98 to aspartate).61,63 This group of autosomal dominant inherited ED disorders are characterized by paronychia, pachyonychia, hyperkeratosis and hyperhydrosis of palms and soles, pilosebaceous cysts, verrucous lesions of the knees, elbows, ankles and buttocks, alopecia and dry hair, natal teeth and corneal defects. JadassohnLewandowsky type is distinct from Jackson-Lawler type by the presence of oral leukoplakia. Beyond these four genetically characterized conditions, EDA, FTA, JadassohnLewandowsky and Jackson-Lawler, there are ten other ED conditions that have specific candidate genes localized to a region of a chromosome. However, the unique mutated gene has not yet been cloned, and in certain cases the chromosomal location is still subject to further confirmation (Table 2.1). Four such conditions are X-linked, including dyskeratosis congenita or Zinsser-Cole-Engman syndrome (OMIM #305000), which is characterized by hypotrichosis, early carious degeneration, dystrophic nails, paronychia, hyperkeratosis,

22

Molecular Basis of Epithelial Appendage Morphogenesis

hyperpigmentation and multiple skin lesions. Leucoplakia is common on various mucosal linings; obliteration of lacrimal ducts and hematologic disturbances such as anemia are all frequent. Incongenitia pigmenti (IP) I and II (OMIM #308300 and #308310, respectively) are both severe X-linked dominant conditions with male lethality. The classical clinical phenotype includes cutaneous vesicular inflammatory eruptions which heal to leave verrucous lesions and “marble-pattern” pigmentation. Defects in hair, nail, teeth and eye and mental retardation are common. The two forms of IPs are distinguishable by genetic markers (i.e., Xp11.21 versus Xq28) and occurrence (i.e., sporadic versus familial). The phenotypic manifestations appear to be similar. The mouse models tattered and striated are suggested to reflect the same genetic disorders as IPI and IPII, respectively, based on their characteristic skin lesions, but molecular confirmation is not as yet available.64-66 Finally, another X-linked dominant syndrome is focal dermal hypoplasia, or Goltz syndrome (OMIM #305600), which illustrates defects in all four cardinal signs, including multiple cutaneous defects such as absence of skin patches, thinness, hyperpigmentation, herniation, hyperkaratosis and papillomatous lesions. Frequently, it is associated with limb defects, eye defects, microcephaly and mental retardation. Three additional conditions of autosomal dominant transmission have been extensively studied. One such condition is hydrotic ectodermal dysplasia, also known as Clouston syndrome (OMIM #129500). Affected individuals present hypotrichosis, hypodontia, nail defects and normal functional sweat glands. Other manifestations include skin hyperpigmentation, palmar and plantar dyskeratosis, strabismus and clubbing of fingers. The mouse model naked has been proposed to be homologous.67 Trichorhinophalangeal syndrome (OMIM #190350) literally implies defects of the hair, nose and digits, but of diverse severity and type. Defects in teeth, nails, eyes and skeleton with mental retardation and short stature have been described. As the name suggests, ectrodactyly, ectodermal dysplasia and cleft lip/ palate (EEC) syndrome (OMIM #129900) encompasses defects in three major tissues, and can also include a plethora of defects in hearing, vision, psychomotor capabilities and urogenital system. Three other ED conditions represent autosomal recessive modes of transmission, and contain skeletal defects in their clinical phenotype. Ellis-Van Creveld syndrome (EvC; OMIM #225500) is characterized by hypotrichosis, hypodontia, natal teeth, nail dysplasia, craniofacial anomalies, acromelic dwarfism, bilateral postaxial polydactyly and congenital heart diseases. Interestingly, the suspected gene has been mapped to 4p16 where the genes for Msx-1 (4p16.1) and FGFR3 (4p16.3) have been mapped. Mutations in these two genes are found in patients with adontia and achondroplasia, respectively, phenotypes shared by EvC syndrome.68 Patients with metaphyseal chondrodysplasia (OMIM #250250) exhibit shortlimbed dwarfism, fine, sparse and hypochromic hair, anemia and immunodeficiencies, rendering the individuals more highly susceptible to chickenpox. Rothmond-Thomson syndrome (OMIN #268400) is characterized by defects in hair, nail and teeth, but normal sweat glands. Diagnostic features include telangiectasia, poikiloderma, hyperkeratosis, cataract, craniofacial skeletal anomalies, short stature, short digit and hypogonadism. Significantly, EDs have a broad spectrum of clinical phenotypes and classifications that lead to confusion among basic scientists and clinicians alike. However, from a current developmental biology perspective, it is not surprising that defects primarily of the ectoderm, which by itself gives rise to numerous tissues and organs, can also involve diverse nonectodermal phenotypes. Embryonic processes such as cell migration, apoptosis, epithelial-mesenchymal transdifferentiation, epithelial-mesenchymal interactions and tissue induction can account for the plethora of cells, tissues and organs involved in EDs. Cell-cell and cell-extracellular matrix interactions at the molecular level, therefore, are the key events in governing normal differentiation throughout evolution.

Ectodermal Dysplasia

23

Hierarchical Regulation and Feedback Loops in Epithelial-Mesenchymal Interactions The cloned or isolated genes associated with EDs still only account for a minority of cases of ED. If we consider the molecular and genetic interactions of genes associated with EDs such as Msx-1 and EDA, as well as the signal transduction pathways that regulate epithelial-mesenchymal interactions during development, we can identify candidate pathways of genes that may account for many of the other cases of ED. The exchange of signals between epithelial and adjacent mesenchymal cells is fundamental to coordinating the activities of two cell types during the formation of complex structures in embryonic development. Such examples of epithelial-mesenchymal interactions in determining the subsequent patterning signals have been demonstrated in many tissues: glandular branching morphogenesis (e.g., lungs, mammary glands, glands of the skin, kidneys), hair follicles, tooth, and limb (for reviews, see refs. 69-73). Recent progress to define the developmental dialog between epithelial and mesenchymal cells has evolved from fundamental studies of the developing fruit fly, and of vertebrate tooth and limb. We will discuss these in greater detail as illustrations of functional conservation of molecular events during evolution. Numerous signaling molecules have been identified in developing tooth and limb morphogenesis, such as growth factors and their cognate receptors (e.g., TGFα/EGF family, TGF-β family, FGF family, retinoids, Sonic and Indian hedgehog, Wnt7a), transcription factors (e.g., Hoxs, Msx-1, Msx-2, Evx, Dlx, Lef1), extracellular matrix molecules and their receptors (e.g., integrins, tenascin, collagens, proteoglycans) (for recent reviews, see refs. 70, 73-76). From these analyses, a small group of molecules have been identified as evolutionarily conserved in a series of feedback loops that coordinate epithelial-mesenchymal interactions. These molecules appear to be functionally conserved from Drosophila to human, and are essential for the patterning of limbs, teeth and likely other vertebrate structures. These molecules include members of the families of Wnt, hedgehog, transforming growth factor beta, fibroblast growth factors, and homeobox containing transcription factors.

Epithelial/Mesenchymal Interactions in Drosophila The relative simplicity of the developmental program of Drosophila, combined with well developed genetic tools for research in this organism, has allowed the identification and characterization of many genes that are responsible for the patterning of the body plan, morphogenesis and cytodifferentiation. The wingless (Wg) gene in Drosophila encodes the prototype of a family of secreted signaling glycoproteins that associates with the cell surface and extracellular matrix. The Wg gene product functions in a number of morphogenetic events including patterning of the wing imaginal disc, the determination of dorso-ventral and proximo-distal axes of the limbs, defining segment polarity of the embryonic epidermis, and the formation of constrictions in the midgut (for reviews, see refs. 77-79). The evolutionary conservation of mechanisms of signaling are perhaps best depicted in the midgut, where Wg participates in mesenchymal to epithelial communication. Wg expressed in mesoderm induces the expression of the homeobox protein labial in the underlying endoderm.81 Wg does not act alone in this induction, but requires signaling by decapentaplegic (Dpp), a member of the TGF-β family and secreted by another distinct domain of mesodermal cells.82 These events provide a morphoregulatory function in the developing fly embryo that leads to the formation of the second midgut constriction. At another site, in the epidermis, Wg regulates the patterning and established polarity in the individual segments that make up the fly body plan. Wg expression from certain epidermal cells signals neighboring cells to maintain their expression of the homeobox transcription factor engrailed (En). En activates hedgehog (hh) transcription.81 After its secretion, hh undergoes autocatalytic cleavage

24

Molecular Basis of Epithelial Appendage Morphogenesis

and coupling onto cholesterol, which limits the distance that it can diffuse, an important feature in maintaining the precise cell-to-cell communication necessary for this developmental process.83 Hh feeds back through patched (ptc) and smoothened to activate the expression of Wg, thus closing the activation feedback loop to define segment polarity in the embryo.84,85 This type of signaling loop allows for short range signals to be propagated through neighboring cells or tissues, and tightly regulates the expression level of potent morphogens, a theme maintained throughout evolution in vertebrates and certain invertebrates. Genes with significant homology to Wg, Dpp and the homeobox family are conserved in vertebrates in regulating development and morphogenesis, especially through epithelialmesenchymal signaling. Through evolution these signaling elements have adapted into combinations suitable for their specific tasks in vertebrate development; significantly their order of interaction has been altered within a combinatorial process. However, an important aspect of the interaction of these signaling pathways preserved through evolution is the establishment of feedback loops that maintain the expression of key signaling molecules and synchronize the activities of neighboring cell types. The vertebrate limb model The vertebrate limb model is further testament to the importance of epithelial-mesenchymal interactions in the formation of complex epidermal appendage structures. Signaling between adjacent heterotypic cell types establishes and maintains the three axes of pattern formation in the developing limb (Fig. 2.1). Each of the three axes of patterning has been attributed to the action of a specific signaling molecule: 1. the proximo-distal (P-D) axis (the shoulder to the finger tips) by FGF-2 and/or FGF-4; 2. the antero-posterior (A-P) axis (the thumb to the little finger) by Sonic hedgehog (Shh); and 3. the dorso-ventral (D-V) axis (the back of the hand to the palm) by Wnt7a. These signaling molecules can substitute for tissue compartments that establish the three axes (see Fig. 2.1A); the apical ectodermal ridge (AER) for P-D axis, the zone of polarizing activity (ZPA) for the D-V axis, and the dorsal ectoderm for the D-V axis. These tissue regions arise in concert and are maintained interdependently (Fig. 2.1B; for reviews, see refs. 75, 76, 86). The first morphological evidence of limb formation is a thickening of a small region of the trunk ectoderm. The earliest known signal involved in limb bud development is FGF-10, expressed in the prospective limb mesoderm and serving as an endogenous limb initiator.87 Expression is maintained in the limb mesenchyme, which suggests a secondary role in limb outgrowth. Ectopic expression of FGF-10 can induce FGF-8 expression and the formation of additional limbs. FGF-8 is important in limb bud development, expressed by cells in the intermediate mesoderm adjacent to somites 15-20.88,89 A physical barrier between the intermediate and lateral plate mesoderm, presumably blocking FGF-8 signaling, will result in absence of limb buds.90 Implantation of FGF-2, -4 or -8-soaked beads in chicken embryos results in the formation of extra limbs.91,92 Subsequent to limb bud initiation, there is extensive proliferation of the lateral plate mesoderm underlying this thickened ectoderm, which results in the outgrowth and formation of the limb bud. The now thickened pseudostratifiedepithelial ectoderm at the distal tip of the developing limb bud forms a ridge that runs along the antero-posterior axis, the apical ectodermal ridge (AER; Fig. 2.1A). The AER is an important source of signals for the underlying mesenchyme, known as the progress zone (Fig. 2.1A). In addition to initiating limb bud outgrowth, FGFs are also essential for patterning the P-D axis. FGF-2 and/or FGF-8 secreted by the AER promotes proliferation and restricts differentiation of the mesenchymal cells in the progress zone.93,94 The fate of mes-

Ectodermal Dysplasia

25

Fig. 2.1. (A)The developing A limb bud has three axes of patterning: proximo-distal (shoulder to finger tips), antero-posterior (thumb to little finger), and dorsoventral (back of hand to palm). Proximo-distal patterning is established by signals from the apical ectodermal ridge, a thickened region of epithelium. Antero-posterior patterning is initiated and maintained by the zone of polarizing activity (ZPA), a specialized region of mesoderm. The dorsal ectoderm patterns the dorso-ventral axis of B the limb. (B) Specific signaling molecules are responsible for establishing the pattern of the limb bud. The developmental activity of the epithelium and the mesenchyme is coordinated by a feedback loop. FGF-4 can supply the function of the apical ectodermal ridge98 which, along with Wnt7a from the dorsal ectoderm,99 acts on the posterior mesenchyme to induce and maintain the function of the ZPA.100 In addition, Wnt7a is required for maintaining dorso-ventral pattern.101 Sonic hedgehog provides the polarizing activity of the ZPA96 and induces BMP-2 expression in the adjacent mesenchyme.97,102 BMP-2 then feeds back on the epithelium to maintain FGF-4 expression in the AER.103

enchymal cells is thought to be determined by the length of time they spend in the progress zone,95 perhaps by exposure to a gradient concentration of FGFs. As the limb bud extends, mesenchymal cells in the proximal portion of the progress zone encounter progressively lower concentrations of FGF from the AER. The mesenchymal cells eventually drop out of the advancing progress zone and begin differentiating into skeletal elements of the arm, forearm and hand. As the AER patterns the P-D axis, the ZPA patterns the A-P axis. The ZPA is a specialized region of mesoderm at the posterior margin of the limb bud, just proximal to the progress zone (Fig. 2.1A). When the ZPA from one limb bud is transplanted to an anterior position of another limb bud, mirror image axis duplication of the distal structures of the limb results, with the formation of additional digits.96,97 Shh was able to substitute for the ZPA in normal and extra A-P axis formation.98 Shh expression is induced by FGF-4 from

26

Molecular Basis of Epithelial Appendage Morphogenesis

the AER (Fig. 2.1B; refs. 99, 100), but this induction is contingent upon a signal from the dorsal ectoderm. Wnt7a, a member of the Wg family, is expressed exclusively in the dorsal ectoderm101 and can substitute for the ectoderm in maintaining the expression of Shh (Fig. 2.1B; ref. 102). In support of this, the phenotype of a Wnt7a gene knockout in mice is the loss of D-V axis, with foot pads developing on the dorsal surface of the paw, and the loss of posterior structures such as the ulna and digit 5.103 Once Shh expression in the ZPA is established, it initiates a feed-back loop to maintain its own induction from the AER (Fig. 2.1B). This loop proceeds from the ZPA to the progress zone, where cells express bone morphogenetic protein-2 (BMP-2). Ectopic placement of the ZPA or ectopic expression of Shh results in the induction of BMP-2 in the mesoderm.99,104 BMP-2 is a TGF-β family member and a vertebrate homologue of Dpp. The BMP-2 signal induces FGF-4 expression in the AER,105 thus closing the loop to Shh (Fig. 2.1B). Two additional molecules are critical: Msx-1 and Msx-2. Msx-1 and 2 are homeobox transcription factors whose expression patterns suggest their function in epithelial-mesenchymal signaling, regulating the morphogenesis and differentiation not only of limb bud but also of tooth bud (discussed in the next section), and other tissues (for review, see ref. 106). These transcription factors are proteins of approximately 260 amino acids, with a centrally located 60 amino acid DNA binding homeodomain. There are 2 amino acid differences between the homeodomain of Msx-1 and 2; so it is likely that these transcription factors regulate many of the same downstream genes. Both of these Msx family members are localized in the mesenchyme of the progress zone in the developing limb bud, but Msx-2 is also expressed in the AER.106-109 At these instructive sites, the paracrine action of BMPs and FGFs may regulate their expression. The expression of Msx-1 in the developing limb bud can be maintained by the application of beads soaked with either BMP-4 or FGF-2 in the absence of the AER.110,111 Furthermore, BMP-4 will induce the expression of both Msx-1 and Msx-2 in dental mesenchyme.72,112 The overlapping expression pattern and regulated expression of Msx-1 and 2 suggests an overlap in function, which is also supported by their molecular architecture and the phenotype of their mutants (discussed in next section). Later during limb development, Msx-2 expression is more restrictively associated with regions where apoptosis is prevalent.113 Interestingly, a causal link has been proposed among BMP-4 expression, Msx-2 expression and apoptosis in rhombomeres 3 and 5 of the developing hindbrain.114 Exogenous BMP-4 treatment of rhombomere cells in culture results in Msx-2 expression and cell death. This process is thought to eliminate the number of neural crest cells derived from the odd-numbered rhombomeres.

Tooth as a Model The developing tooth bud is another well characterized model for defining the molecular mechanisms required for reciprocal signaling during epithelial-mesenchymal interactions in morphogenesis and cytodifferentiation.115-117 Prior to mouse embryonic day 12, dental epithelium can induce tooth bud formation in nondental, neural crest derived mesenchyme. However, after day 12, this inductive potential shifts so that the dental mesenchyme is inductive to the epithelium.118,119 This shift in dominance temporally corresponds with a change in the expression pattern of BMP-4 from the epithelium to the condensing mesenchyme.112 BMP-2 is also expressed in the early dental epithelium and shifts to the mesenchymal dental papilla at the late cap stage.112 BMPs provide key signals for the combinatorial process between epithelium and mesenchyme. At the initiation stage (E11.5), BMP-4 in the epithelium either activates or maintains Msx-1 expression in the mesenchyme. A bead soaked with BMP-4 can substitute for dental epithelium in inducing dental mesenchyme differentiation, including the expression of Msx-1 and Msx-2.70 However, in the Msx-1 null mutation transgenic mouse model, there is a decrease of expression of BMP-4, the

Ectodermal Dysplasia

27

Fig. 2.2. Several regulatory genes have been associated with ectodermal dysplasia syndromes in humans and in mouse models. Based on genetic or biochemical data, these genes can be arranged into a pathway that may regulate the patterning and differentiation of specific cell types at sites of epithelial-mesenchymal interaction. In the tooth bud mesenchyme, BMP-4 induces the expression of Msx-1. 110,117 In turn, Msx-1 feeds back to regulate the expression of BMP-4, which is reduced in the Msx-1 null mutant mouse.117 However, Lef1 expression is induced by BMP-4 in both normal and Msx-1 null mutant dental mesenchyme.117,118 Tooth bud development arrests at the same stage in Msx-1 and Lef1 null mutant mice, and the Msx-1 mutation causes a decrease in Lef1 expression.117,119 Lef1 is expressed in developing tooth buds, hair follicles, and other sites of epithelial-mesenchymal interaction,119,141 and a putative Lef1 binding site is found in the upstream, noncoding region of the EDA gene.35 EDA/Ta is connected with the EGF receptor signaling pathway, since EGF treatment rescues developmental defects in the Tabby mouse,125,130 and EGF receptor expression is decreased in EDA patients.131

transcription factor Lef1 and the extracellular matrix component syndecan-1 in dental mesenchyme.120 A bead soaked in BMP-4 can induce expression of BMP-4 and Lef1 in wild type mesenchyme, but only Lef1 in the Msx-1 mutant mesenchyme.120,121 Furthermore, BMP-4 beads can induce the expression of Msx-1 in Lef1 null mutant mesenchyme.121 Taken together, these data suggest that epithelial BMP-4 regulates the expression of mesenchymal BMP through Msx-1, and that Lef1 expression is regulated by the mesenchymal BMP (see Fig. 2.2). Lef1 is required for the development of teeth and whiskers in transgenic mice.122 However transplantation experiments between normal and Lef1 null mutant mouse tissues demonstrated that Lef1 expression in the dental mesenchyme is not required for tooth development, and Lef1 expression in the dental epithelium is only transiently required.121 This suggests that Lef1 in the epithelium regulates the expression of some extracellular signaling factor that communicates with the underlying mesenchyme to overcome a hurdle in tooth development. The identity of this signaling factor awaits discovery. Limb bud and tooth bud patterning are both dependent on epithelial-mesenchymal communication involving some of the same signaling molecules. Msx-2 is expressed in the mesenchyme of the dental papilla and in the epithelium, especially in the enamel knot, which is found to have a role in patterning the tooth organ, a function comparable to that of the AER of the developing limb bud.123 Interestingly, Shh is also expressed in the enamel knot, likely contributing to tooth bud patterning.124 Koyama et al125 demonstrated that transplanting the enamel knot from a mouse embryo to an anterior position in a chicken limb

Molecular Basis of Epithelial Appendage Morphogenesis

28

bud induced supernumerary digits. The enamel knot in this experiment exhibited limb polarizing activity like that of the ZPA, likely due to the expression of Shh. These data suggest that two of the three axes of pattern formation in the tooth bud are initiated or maintained by the enamel knot. Further studies will be necessary to understand: 1. how this one structure provides these two instructions; and 2. what the structure or molecule is that provides the signal for the third axis of patterning, buccal vs. lingual.

Animal Models for Ectodermal Dysplasias Whereas genetics from Drosophila and other invertebrates provide insights into the range of signaling candidate molecules that may participate in epithelial-mesenchymal interactions, mouse models for human genetic diseases often mimic the same or comparable clinical phenotypes. Because of similarities in genome organization, mouse models are invaluable in the cloning of human disease genes such as EDA. Furthermore, mice can provide an appropriate testing model for potential clinical therapeutics. Several spontaneously arising mouse mutants have been described that mimic ectodermal dysplasias (see Table 2.2), the best characterized being Tabby (Ta; ref. 126). The Ta gene is the mouse homolog of EDA.127 Like EDA, Ta is an X-linked gene with males more affected than females. The normal mouse coat contains 4 types of hairs, but Ta mice have only one, abnormal hair type, which is present in reduced numbers, and their tails are bald. Female heterozygotes have an abnormal pattern of coat color, with alternating bands of light and dark, and thus they were given the name Tabby. The light bands are normal coat and the dark bands are abnormal, arising from the random pattern of X-chromosome inactivation. In addition to the abnormal coat texture of Ta mice, there are deleterious effects on the eyelids, teeth, sweat and mucous glands that in many ways parallel defects seen in human EDA. Ta mice exhibit delayed postnatal eyelid opening and reduced aperture of the eyelids.128 They develop ulcerations of the cornea. The incisors and molars are delayed in eruption, reduced in size and sometimes maloccluded, but usually all teeth are present.129,130 Mice normally have sweat glands only on their foot pads, but these are absent in Ta mice.131 Ta mice exhibit a high frequency of respiratory tract infections that are often lethal. Body hairs are easily dislodged during grooming and become trapped in the air passages, contrib-

Table 2.2. Mouse models for ectodermal dysplasias Gene or Mutant Name *

Chromosome

Relation to ED Number

Tabby (Ta) Crinkled (Cr) Downless (Dl) Sleek (Slk) TGF alpha,waved-1

X 13 10 10 6

EGF receptor,waved-2

11

Lef1

3

likely equivalent to EDA identical phenotype to Ta phenotype closely resembles Ta probably allelic to Dl wavy hair and whiskers, eye abnormalities like Ta wavy hair and whiskers, eye abnormalities like Ta failure to develop teeth, hair, whiskers, and mammary glands

* Mouse Genome Database, 1996 (MGD; ref. 135).

Ectodermal Dysplasia

29

uting to increased infections. The abnormal phenotype of the Ta mouse demonstrates striking resemblance to the human EDA clinical phenotype, something that is not always true of mutations in homologous genes of different organisms. This could make the Ta mouse instrumental in the discovery of therapeutics for the management of human EDs. Evidence suggests that the Ta gene participates in a signal transduction pathway involving epidermal growth factor (EGF) (see Fig. 2.2). Cohen and co-workers in 1962 described the effects of treating normal neonatal mice with exogenous EGF.132 The results were precocious eyelid opening and precocious tooth eruption, phenotypically opposite of the Ta mouse. This observation suggests tests to determine the effects of EGF treatment on Ta. Injections of EGF into Ta neonates rescued the delay in eyelid opening and incisor tooth eruption, and promoted the development of functional sweat glands.128,133 Further support for the involvement of Ta and EDA in the EGF signaling pathway came from studies of EGF receptor levels. Fibroblasts from HED patients were found to have a 2- to 8-fold reduction in the binding of EGF, and reduced receptor levels were confirmed at both the mRNA and protein levels.134 Similarly, EGF receptor expression was low in fibroblasts and liver membranes of Ta mice.134 Further characterization of the EDA protein will be necessary for understanding of its function in relation to the EGF signaling pathway. The Ta/EDA gene is but one step in a pathway regulating tooth development and other epidermal derivatives of the skin. Other genetic mutations affecting the same pathway can give rise to the same phenotype. Crinkled (Cr), Downless (Dl) and Sleek (Slk) are other strains that present a phenotype indistinguishable from Ta. None of these genes has yet been cloned, but certainly represent opportunities to understand and treat EDs. Cr is a recessive mutation found on chromosome 13. Despite the striking similarity of homozygous Cr with hemizygous Ta, fitting Cr into the Ta pathway presents a bit of a puzzle. Cr mice have been characterized as deficient in copper metabolism; copper levels are low in tissues of Cr mice, and the level of copper dependent superoxide dismutase (SOD) activity is also low.135,136 Normal development of skin and hair, as well as SOD level in Cr mice is restored by prenatal supplementation with copper.136,137 Dl and Slk are both on chromosome 10 and may be allelic. Dl is recessive and Slk is dominant, and their affected phenotype is similar to that of Cr. Shawlot and co-workers described a mouse strain with a transgene inserted into the Dl locus.138 This provided probes that are close to or within the Dl gene and should facilitate cloning and characterization of this potentially important gene. Another ED-like syndrome is found in the waved-1 (Wa-1) and Wa-2 mouse strains. These animals exhibit waviness of whiskers and fur, malformations within the hair follicles, open eyelids at birth, reduced eyeball size, inflammation and scarring of the corneas (like Ta), lens and retinal defects. Wa-1 represents a mutation in the TGF alpha gene, and the TGF alpha null mutation gives the same abnormal phenotype.139,140 TGF alpha is another ligand of the EGF receptor. Wa-2 is a mutation in the kinase domain of the EGF receptor causing a greater than 90% loss of tyrosine kinase activity.141 This is further evidence for multiple mutations in the EGF receptor signaling pathway giving rise to EDs. Perturbation of EGF receptor signaling in the developing tooth organ and limb bud produce dysmorphogenesis.142-144 Consistent with the hypothesis that EDA expression is regulated by the transcription factor Lef1, changes in the expression pattern of Lef1 in mice generates abnormal phenotypes similar to EDs. Lef1 is expressed in developing hair follicles, beginning even before the morphological initiation, and continuing throughout hair maturation, with high expression in the dermal papilla. Lef1 expression is also identified in the developing tooth. During the odontogenic placode stage, Lef1 is expressed in the dental epithelium;145 however, beginning with the bud stage and up to the cap stage, expression is co-localized to both dental epithelium and adjacent dental mesenchyme.121,122 Lef1 is expressed in a number of

30

Molecular Basis of Epithelial Appendage Morphogenesis

cell lineages including neural crest cells, mesencephalon, otic placode, branchial arches, nasal process, pituitary gland, choroid plexus, limb bud, lung, urogenital system, kidney and thymus.122,145 Overexpression of Lef1 using the keratin K14 promoter results in disruption in hair, whisker and tooth formation.49 The targeted disruption of the Lef1 gene on chromosome 3 resulted in postnatal lethality with adontia, hypoplastic mammary glands and hypotrichosis.122 This ED-like abnormal phenotype suggests that Lef1 and EDA may be two consecutive steps in a signaling pathway regulating epithelial-mesenchymal interactions (see Fig. 2.2). The mapping of this signaling pathway will likely lead to an understanding of the genetic mechanisms underlying the array of ED syndromes. The Msx-1 and Lef1 null mutant mice each exhibit tooth bud developmental arrest at the same stage, suggesting that they reside in the same genetic pathway.25,122 Lef1 expression is reduced in the Msx-1 mutant dental mesenchyme120 while Msx-1 expression is maintained in the dental mesenchyme of Lef1 mutant mice.121 This suggests that Msx-1 is epistatically higher than Lef1 in the same genetic pathway (see Fig. 2.2).

Prospectus When a stem cell decides to leave its self renewal phase to enter into its differentiative phase, it has to go through a series of decisions, which are subject to influence from its near and far neighbors. It initiates a developmental program in which it draws from its intrinsic genetic information and environmental milieu to activate transcription of a set of genes that will progressively restrict the fate of the cell. In this scenario, the cell will undergo determination, commitment and terminal differentiation, coinciding with a shift from totipotency to monopotency; and a shift from a proliferative to a postmitotic state. The intrinsic elements are modifiable by extrinsic factors such as cell-cell communication through direct cell contact and by long-range diffusable factors or short range extracellular matrix binding factors. When each cell within a group of cells undergoes its own developmental program, but the cells interact to regulate one another, then there is pattern formation. This is likely to be the logic for epithelial-mesenchymal interactions. Ectodermal dysplasia is a particularly suitable model for illustrating a complex human disease with clinical phenotypes, that cannot, however, begin to suggest the etiology of this large set of disorders. It would now seem imperative to explore and understand the developmental, cellular, molecular and evolutionary biology behind the clinical phenotypes. There are so many opportunities to examine ectodermal defects as they relate to heterotypic tissue interactions, the time and position for these instructive interactions and the various molecular feedback mechanisms that characterize epithelial appendage morphogenesis. Many questions need to be addressed for ectodermal dysplasias and for the broader issues of epithelial-mesenchymal interactions: Does EGF receptor mediate the function of EDA; if so, how? What might be the members of the TGFα/EGF family of ligands that acts on the EGF receptor in EDA? How does EDA fit into the regulatory feedback loops of tissue patterning? Are mutations in genes implicated in epithelial-mesenchymal interactions associated with EDs? What are the compensatory mechanisms for disruptions to epithelial-mesenchymal interactions? What can we learn from nature’s experiment of mutant mouse strains?

Acknowledgments We wish to dedicate this manuscript to Mary Kay Richter and the many heroic families and their children who champion the research agenda of the National Ectodermal Dysplasia Foundation.

Ectodermal Dysplasia

31

References 1. Darwin C. The Variation of Animals and Plants under Domestication. 2nd edition. London: John Murray,1875:319. 2. Freire-Maia N, Pinheiro M. Ectodermal Dysplasias: A Clinical and Genetic Study. 1984; Alan R. Liss, Inc., New York. 3. Levine M, Rubin GM, Tjian R. Human DNA sequences homologous to a protein coding region conserved between homeotic genes of Drosophila. Cell 1984; 38(3):667-673. 4. McGinnis W, Garber RL, Wirz J et al. A homologous protein-coding sequence in Drosophila homeotic genes and its conservation in other metazoans. Cell 1984; 37(2):403-408. 5. McGinnis W, Levine MS, Hafen E et al. A conserved DNA sequence in homoeotic genes of the Drosophila Antennapedia and bithorax complexes. Nature 1984; 308(5958):428-433. 6. McClain DA, Edelman GM. A neural cell adhesion molecule from human brain. Proc Nat Acad Sci USA 1982; 79(20):6380-6384. 7. Thiery JP, Duband JL, Rutishauser U et al. Cell adhesion molecules in early chicken embryogenesis. Proc Nat Acad Sci USA 1982; 79(21):6737-6741. 8. Hoffman S, Edelman GM. The mechanism of binding of neural cell adhesion molecules. Adv Exp Med Biol 1984; 181:147-160. 9. Kauffman SA. Pattern formation in the Drosophila embryo. Philosophical Transactions of the Royal Society of London. Series B: Biological Sciences 1981; 295(1078):567-594. 10. Slavkin HC. Combinatorial process for extracellular matrix influences on gene expression: A hypothesis. J Cranio Gen Dev Biol 1982; 2(2):179-189. 11. Gray A, Dull TJ, Ullrich A. Nucleotide sequence of epidermal growth factor cDNA predicts a 128,000-molecular weight protein precursor. Nature 1983; 303(5919): 722-725. 12. Ullrich A, Gray A, Berman C et al. Human beta-nerve growth factor gene sequence highly homologous to that of mouse. Nature 1983; 303(5920):821-825. 13. Isackson PJ, Ullrich A, Bradshaw RA. Mouse 7S nerve growth factor: Complete sequence of a cDNA coding for the alpha-subunit precursor and its relationship to serine proteases. Biochemistry 1984; 23(25):5997-6002. 14. Scott J, Urdea M, Quiroga M et al. Structure of a mouse submaxillary messenger RNA encoding epidermal growth factor and seven related proteins. Science 1983; 221: 236-240. 15. Ullrich A, Coussens L, Hayflick JS et al. Human epidermal growth factor receptor cDNA sequence and aberrant expression of the amplified gene in A431 epidermoid carcinoma cells. Nature 1984; 309(5967):418-425. 16. Ullrich A, Gray A, Wood WI et al. Isolation of a cDNA clone coding for the gammasubunit of mouse nerve growth factor using a high-stringency selection procedure. DNA 1984; 3(5):387-392. 17. Johnston MC, Bronsky PT. Prenatal craniofacial development: New insights on normal and abnormal mechanisms. Crit Rev Oral Biol Med 1995; 6:368-422. 18. Slavkin HC. Molecular biology experimental strategies for craniofacial-oral-dental dysmorphology. Connect Tissue Res 1995; 32:233-239. 19. Slavkin HC. Meeting the challenges of craniofacial-oral-dental birth defects. J Am Dent Assoc 1996; 127:681-682. 20. Chuong CM, Widelitz RB, Ting-Berreth S et al. Early events during avian skin appendage regeneration: Dependence on epithelial-mesenchymal interaction and order of molecular reappearance. J Invest Dermatol 1996; 107:639-646. 21. Davidson D. The function and evolution of Msx genes: Pointers and paradoxes. Trends Genet 1995; 11(10):405-411. 22. Maas R, Chen YP, Bei M et al. The role of Msx genes in mammalian development. Ann N Y Acad Sci 1996; 785:71-181. 23. Ignelzi MA Jr, Liu YH, Maxson RE Jr et al. Genetically engineered mice: Tools to understand craniofacial development. Crit Rev Oral Biol Med 1995; 6:181-201. 24. Liu YH, Kundu R, Wu L et al. Premature suture closure and ectopic cranial bone in mice expressing Msx2 transgenes in the developing skull. Proc Natl Acad Sci U S A 1995; 92(13):6137-6141.

32

Molecular Basis of Epithelial Appendage Morphogenesis

25. Satokata I, Maas R. Msx1 deficient mice exhibit cleft palate and abnormalities of craniofacial and tooth development. Nat Genet 1994; 6(4):348-356. 26. Bronner-Fraser M. Origins and developmental potential of the neural crest. Exp Cell Res 1995; 218:405-417. 27. Hay ED. An overview of epithelio-mesenchymal transformation. Acta Anat 1995; 154:8-20. 28. Shuler CF. Programmed cell death and cell transformation in craniofacial development. Crit Rev Oral Biol Med 1995; 6:202-217. 29. Viebahn C. Epithelio-mesenchymal transformation during formation of the mesoderm in the mammalian embryo. Acta Anat 1995; 154:79-97. 30. McKusick VA, Amberger JS, Francomano CA. Progress in medical genetics: map-based gene discovery and the molecular pathology of skeletal dysplasias. Am J Med Genet 1996; 63:98-105. 31. Lewanda AF, Meyers GA, Jabs EW. Craniosynostosis and skeletal dysplasias: Fibroblast growth factor receptor defects. Proc Assoc Am Physicians 1996; 108:19-24. 32. Malcolm S, Reardon W. Fibroblast growth factor receptor-2 mutations in craniosynostosis. Ann N Y Acad Sci 1996; 785:164-170. 33. Webster MK, Donoghue DJ. FGFR activation in skeletal disorders: Too much of a good thing. Trends Genet 1997;13:178-82. 34. Pinheiro M, Freire-Maia N. Ectodermal dysplasias: A clinical classification and causal review. Am J Med Gen 1994; 53:153-162. 35. Weech AA Heriditary ectodermal dysplasia (congenital ectodermal defects). A report of two cases. Am J Dis Child 1929; 37:766-790. 36. Kere J, Srivastava AK, Montonen O et al. X-linked anhydrotic (hypohydrotic) ectodermal dysplasia is caused by mutation in a novel transmembrane protein. Nat Genet 1996; 13:409-416. 37. Chautard-Freire-Maia EA, Primo-Parmo SL, Pinheiro M et al. Further evidence against linkage between Christ-Siemens-Touraine (CST) and Xg loci. Hum Genet 1981; 57:205-206. 38. Kolvraa S, Kruse TA, Jensen PKA et al. Close linkage between X-linked ectodermal dysplasia and a cloned DNA sequence detecting a two allele restriction fragment length polymorphism in the region Xp11-q12. Hum Genet 1986; 74:284-287. 39. MacDermot, KD, Winter RM, Malcolm S. Gene localization of X-linked hypohydrotic ectodermal dysplasia (C-S-T syndrome). Hum Genet 1986; 74:172-173. 40. Clarke A, Sarfarazi M, Thomas NST et al. X-linked hypohydrotic ectodermal dysplasia: DNA probe linkage analysis and gene localization. Hum Genet 1987; 75:378-380. 41. Hanauer A, Alembik Y, Arveiler B et al. Genetic mapping of anhydrotic ectodermal dysplasia: DXS159, a closely linked proximal marker. Hum Genet 1988; 80:177-180. 42. Zonana J, Clarke A, Sarfarazi M et al. X-linked hypohydrotic ectodermal dysplasia: Localization within the region Xq11-21.1 by linkage analysis and implications for carrier detection and prenatal diagnosis. Am J Hum Genet 1988; 43:75-85. 43. Turleau C, Niaudet P, Cabanis MO et al. X-linked hypohydrotic ectodermal dysplasia and t(X;12) in a female. Clin Genet 1989; 35:462-466. 44. MacDermot, KD, Hulten M. Female with hypohydrotic ectodermal dysplasia and de novo (X;9) translocation: Clinical documentation of the AnLy cell line case. Hum Genet 1990; 84:577-579. 45. Limon J, Filipiuk J, Nedoszytko B et al. X-linked anhydrotic ectodermal dysplasia and de novo t(X;1) in a female. Hum Genet 1991; 87:338-340. 46. Zonana J, Jones M, Browne D et al. High-resolution mapping of the X-linked hypohydrotic ectodermal dysplasia (EDA) locus. Am J Hum Genet 1992; 51:1036-1046. 47. Zonana J, Gault J, Davies KJP et al. Detection of a molecular deletion at the DXS732 locus in a patient with X-linked hypohydrotic ectodermal dysplasia (EDA), with the identification of a unique junctional fragment. Am J Hum Genet 1993; 52:78-84. 48. Rogers GE, Powell BC. Organization and expression of hair follicle genes. J Invest Dermatol 1993; 101:50S-55S. 49. Zhou P, Byrne C, Jacobs J et al. Lymphoid enhancer factor 1 directs hair follicle patterning and epithelial cell fate. Genes Dev 1995; 9:700-713.

Ectodermal Dysplasia

33

50. Graber LW. Congenital absence of teeth: A review with emphasis on inheritance patterns. J Am Dent Assoc 1978; 96:266-275. 51. Vastardis H, Karimbux N, Guthua SW et al. A human Msx1 homeodomain missense mutation causes selective tooth agenesis. Nat Genet 1996; 13(4):417-421. 52. Lyngstadaas SP, Nordbo H, Gedde-Dahl T et al. On the genetics of hypodontia and microdontia: Synergism or allelism of major genes in a family with six affected members. J Med Genet 1996; 33: 137-142. 53. Hill RE, Jones PF, Rees AR et al. A new family of mouse homeo box-containing genes: Molecular structure, chromosomal location, and develpmental expression of Hox-7.1. Genes Dev 1989; 3:26-37. 54. MacKenzie A, Ferguson MW, Sharpe P. Hox-7 expression during murine craniofacial development. Development 1991; 113:601-611. 55. MacKenzie A, Leeming GL, Jowett AK et al. The homeobox gene Hox 7.1 has specific regional and temporal expression patterns during early murine craniofacial embryogenesis, especially tooth development in vivo and in vitro. Development 1991; 111:269-285. 56. Robert B, Sassoon D, Jacq B et al. Hox-7, a mouse homeobox gene with a novel pattern of expression during embryogenesis. EMBO J 1989; 8:91-100. 57. Graham A, Heyman I, Lumsden A. Even-numbered rhombomeres control the apoptotic elimination of neural crest cells from odd-numbered rhombomeres in the chick hindbrain. Development 1993; 119:233-245. 58. Zhang H, Gezhi H, Wang H et al. Heterodimerization of Msx and Dlx homeoproteins results in functional antagonism. Mol Cell Biol 1997; 17:2920-2932. 59. Jabs EW, Muller U, Li X et al. A mutation in the homeodomain of the human Msx2 gene in a family affected with autosomal dominant craniosynostosis. Cell 1993; 3:443-450. 60. Foerst-Potts L, Sadler TW. Disruption of Msx-1 and Msx-2 reveals roles for these genes in craniofacial, eye, and axial development. Developmental Dynamics 1997; 209:70-84. 61. McLean WHI, Rugg EL, Lunny DP et al. Keratin 16 and keratin 17 mutations cause pachyonychia congenita. Nat Genet 1995; 9:273-278. 62. Bowden PE, Haley JL, Kansky A et al. Mutation of a type II keratin gene (K6a) in pachyonychia congenita. Nat Genet 1995; 10(3):363-365. 63. Smith FJ, Corden LD, Rugg EL et al. Missense mutations in keratin 17 cause either pachyonychia congenita type 2 or a phenotype resembling steatocystoma multiplex. J Invest Dermatol 1997; 108:220-223. 64. Cannizzaro LA, Hecht F. The gene for incontinentia pigmenti maps to band Xp11 with an (X;10)(p11;q22) translocation. Clin Genet 1987; 32(1): 66-69. 65. Levin ML, Chatterjee A, Pragliola A et al. A comparative transcription map of the murine bare patches (Bpa) and striated (Str) critical regions and human Xq28. Genome Res 1996; 6:465-477. 66. Mouse Genome Database (MGD), Mouse Genome Informatics, The Jackson Laboratory, Bar Harbor, Maine. March 1996. World Wide Web URL: http://www.informatics.jax.org. 67. Tenenhouse HS, Gold RJM, Kachra Z et al. Biochemical marker in dominantly inherited ectodermal malformation. Nature 1974; 251:431-432. 68. Polymeropoulos MH, Ide SE, Wright M et al. The gene for the Ellis-van Creveld syndrome is located on chromosome 4p16. Genomics 1996; 35:1-5. 69. Cunha GR. Role of mesenchymal-epithelial interactions in normal and abnormal development of the mammary gland and prostate. Cancer 1994; 74(3):1030-1044. 70. Thesleff I, Vaahtokari A, Kettunen P et al. Epithelial-mesenchymal signaling during tooth development. Connect Tissue Res 1995; 32(1-4):9-15. 71. Hilfer SR. Morphogenesis of the lung: Control of embryonic and fetal branching. Annu Rev Physiol 1996; 58:93-113. 72. Thesleff I, Vaahtokari A, Vainio S et al. Molecular mechanisms of cell and tissue interactions during early tooth development. Anat Rec 1996; 245(2):151-161. 73. Saxén L. Organogenesis of the kidney. Cambridge Univ. Press, Cambridge. 1987. 74. Capecchi MR. Function of homeobox genes in skeletal development. Ann N Y Acad Sci 1996; 785:34-37.

34

Molecular Basis of Epithelial Appendage Morphogenesis

75. Francis-West P, Tickle C. Limb development. Curr Top Microbiol Immunol 1996; 212: 239-259. 76. Niswander L. Growth factor interactions in limb development. Ann N Y Acad Sci 1996; 785: 23-26. 77. Cohen SM, Di Nardo S. Wingless: From embryo to adult. Trends Genet 1993; 9(6):189-192. 78. Klingensmith J, Nusse R. Signaling by wingless in Drosophila. Dev Biol 1994; 166(2):396-414. 79. Siegfried E, Perrimon N. Drosophila wingless: A paradigm for the function and mechanism of Wnt signaling. Bioessays 1994; 16(6):395-404. 80. Immergluck K, Lawrence PA, Bienz M. Induction across germ layers in Drosophila mediated by a genetic cascade. Cell 1990; 62(2):261-268. 81. Panganiban GE, Reuter R, Scott MP et al. A Drosophila growth factor homolog, decapentaplegic, regulates homeotic gene expression within and across germ layers during midgut morphogenesis. Development 1990; 110(4):1041-1050. 82. Dominguez M, Brunner M, Hafen E et al. Sending and receiving the hedgehog signal: Control by the Drosophila Gli protein Cubitus interruptus. Science 1996; 272(5268): 1621-1625. 83. Porter JA, Young KE, Beachy PA. Cholesterol modification of hedgehog signaling proteins in animal development. Science 1996; 274(5285):255-259. 84. Ingham PW, Taylor AM, Nakano Y. Role of the Drosophila patched gene in positional signalling. Nature 1991; 353(6340):184-187. 85. Alcedo J, Ayzenzon M, Von Ohlen T et al. The Drosophila smoothened gene encodes a seven-pass membrane protein, a putative receptor for the hedgehog signal. Cell 1996; 86(2):221-232. 86. Johnson RL, Riddle RD, Tabin CJ. Mechanisms of limb patterning. Curr Opin Genet Dev 1994; 4(4):535-542. 87. Ohuchi H, Nakagawa T, Yamamoto A et al. The mesenchymal factor FGF-10, initiates and maintains the outgrowth of the chick limb bud through interaction with FGF-8 an apical ectodermal factor. Development 1997; 124:2235-2244. 88. Crossley PH, Minowada G, MacArthur CA et al. Roles for FGF-8 in the induction, initiation, and maintenance of chick limb development. Cell 1996; 84(1):127-136. 89. Vogel A, Rodriguez C, Izpisua-Belmonte JC. Involvement of FGF-8 in initiation, outgrowth and patterning of the vertebrate limb. Development 1996; 122(6):1737-1750. 90. Stephens TD, McNulty TR. Evidence for a metameric pattern in the development of the chick humerus. J Embryol Exp Morphol 1981; 61 191-205. 91. Cohn MJ, Izpisua-Belmonte JC, Abud H et al. Fibroblast growth factors induce additional limb development from the flank of chick embryos. Cell 1995; 80(5):739-746. 92. Ohuchi H, Nakagawa T, Yamauchi M et al. An additional limb can be induced from the flank of the chick embryo by FGF-4. Biochem Biophys Res Commun 1995; 209(3):809-816. 93. Crossley PH, Martin GR. The mouse Fgf-8 gene encodes a family of polypeptides and is expressed in regions that direct outgrowth and patterning in the developing embryo. Development 1995; 121(2):439-451. 94. Mahmood R, Bresnick J, Hornbruch A et al. A role for FGF-8 in the initiation and maintenance of vertebrate limb bud outgrowth. Curr Biol 1995; 5(7):797-806. 95. Summerbell D, Lewis JH, Wolpert L. Positional information in chick limb morphogenesis. Nature 1973; 244(5417):492-496. 96. Maccabe AB, Gasseling MT, Saunders J Jr. Spatiotemporal distribution of mechanisms that control outgrowth and anteroposterior polarization of the limb bud in the chick embryo. Mech Ageing Dev 1973; 2(1):1-12. 97. Tickle C, Summerbell D, Wolpert L. Positional signalling and specification of digits in chick limb morphogenesis. Nature 1975; 254(5497):199-202. 98. Riddle RD, Johnson RL, Laufer E et al. Sonic hedgehog mediates the polarizing activity of the ZPA. Cell 1993; 75(7):1401-1416. 99. Laufer E, Nelson CE, Johnson RL et al. Sonic hedgehog and FGF-4 act through a signaling cascade and feedback loop to integrate growth and patterning of the developing limb bud. Cell 1994; 79(6):993-1003.

Ectodermal Dysplasia

35

100. Niswander L, Jeffrey S, Martin GR et al. A positive feedback loop coordinates growth and patterning in the vertebrate limb. Nature 1994; 371(6498):609-612. 101. Dealy CN, Roth A, Ferrari D et al. Wnt-5a and Wnt-7a are expressed in the developing chick limb bud in a manner suggesting roles in pattern formation along the proximodistal and dorsoventral axes. Mech Dev 1993; 43(2-3):175-186. 102. Yang Y, Niswander L. Interaction between the signaling molecules WNT7a and SHH during vertebrate limb development: dorsal signals regulate anteroposterior patterning. Cell 1995; 80(6):939-947. 103. Parr BA, McMahon AP. Dorsalizing signal Wnt-7a required for normal polarity of D-V and A-P axes of mouse limb. Nature 1995; 374(6520):350-353. 104. Francis PH, Richardson MK, Brickell PM et al. Bone morphogenetic proteins and a signalling pathway that controls patterning in the developing chick limb. Development 1994; 120(1):209-218. 105. Duprez DM, Kostakopoulou K, Francis-West PH et al. Activation of FGF-4 and HoxD gene expression by BMP-2 expressing cells in the developing chick limb. Development 1996; 122(6):1821-1828. 106. Davidson DR, Crawley A, Hill RE et al. Position-dependent expression of two related homeobox genes in developing vertebrate limbs. Nature 1991; 352(6334):429-431. 107. Yokouchi Y, Ohsugi K, Sasaki H et al. Chicken homeobox gene Msx-1: Structure, expression in limb buds and effect of retinoic acid. Development 1991; 113(2):431-444. 108. Coelho CN, Sumoy L, Kosher RA et al. GHox-7: A chicken homeobox-containing gene expressed in a fashion consistent with a role in patterning events during embryonic chick limb development. Differentiation 1992; 49(2):85-92. 109. Nohno T, Noji S, Koyama E et al. Differential expression of two msh-related homeobox genes Chox-7 and Chox-8 during chick limb development. Biochem Biophys Res Commun 1992; 182(1):121-128. 110. Vogel A, Roberts-Clarke D, Niswander L. Effect of FGF on gene expression in chick limb bud cells in vivo and in vitro. Dev Biol 1995; 171(2):507-520. 111. Wang Y, Sassoon D. Ectoderm-mesenchyme and mesenchyme-mesenchyme interactions regulate Msx-1 expression and cellular differentiation in the murine limb bud. Dev Biol 1995; 168(2):374-382. 112. Vainio S, Karavanova I, Jowett A et al. Identification of BMP-4 as a signal mediating secondary induction between epithelial and mesenchymal tissues during early tooth development. Cell 1993; 75(1):45-58. 113. Macias D, Ganan Y, Ros MA et al. In vivo inhibition of programmed cell death by local administration of FGF-2 and FGF-4 in the interdigital areas of the embryonic chick leg bud. Anat Embryol 1996; 193(6):533-541. 114. Graham A, Francis-West P, Brickell P et al. The signalling molecule BMP-4 mediates apoptosis in the rhombencephalic neural crest. Nature 1994; 372(6507):684-686. 115. Slavkin H. Tooth development. Adv Dent Res 1995; 9:11. 116. Slavkin HC, Diekwisch T. Evolution in tooth developmental biology of morphology and molecules. Anat Rec 1996; 245:131-150. 117. Maas R, Bei M. The genetic control of early tooth development. Crit Rev Oral Biol Med 1997; 8:4-39. 118. Mina M, Kollar EJ. The induction of odontogenesis in non-dental mesenchyme combined with early murine mandibular arch epithelium. Arch Oral Biol 1987; 32(2):123-127. 119. Lumsden AG. Spatial organization of the epithelium and the role of neural crest cells in the initiation of the mammalian tooth germ. Development 1988; 103:155-169. 120. Chen Y, Bei M, Woo I et al. Msx1 controls inductive signaling in mammalian tooth morphogenesis. Development 1996; 122(10):3035-3044. 121. Kratochwil K, Dull M, Farinas I et al. Lef1 expression is activated by BMP-4 and regulates inductive tissue interactions in tooth and hair development. Genes Dev 1996; 10:1382-1394. 122. van Genderen C, Okamura RM, Farinas I et al. Development of several organs that require inductive epithelial-mesenchymal interactions is impaired in Lef-1-deficient mice. Genes Dev 1994; 8(22):2691-2703.

36

Molecular Basis of Epithelial Appendage Morphogenesis

123. MacKenzie A, Ferguson MW, Sharpe PT. Expression patterns of the homeobox gene, Hox8, in the mouse embryo suggest a role in specifying tooth initiation and shape. Development 1992; 115(2):403-420. 124. Iseki S, Araga A, Ohuchi H et al. Sonic hedgehog is expressed in epithelial cells during development of whisker, hair, and tooth. Biochem Biophys Res Commun 1996; 218(3):688-693. 125. Koyama E, Yamaai T, Iseki S et al, Polarizing activity, Sonic hedgehog, and tooth development in embryonic and postnatal mouse. Dev Dynam 1996; 206:59-72. 126. Falconer DS. Total sex-linkage in the house mouse. Z Indukt Abstamm Vererbungsl 1953; 85:210-219. 127. Pispa J, Srivastava A, Pekkanen M et al. Cloning and expression of Tabby. 13th Int Congress of Dev Biol 1997; in press. 128. Kapalanga J, Blecher SR. Effect of the X-linked gene Tabby (Ta) on eyelid opening and incisor eruption in neonatal mice is opposite to that of epidermal growth factor. Development 1990; 108(2):349-355. 129. Gruneberg H. Genes and genotypes affecting the teeth of the mouse. J Embryol Exp Morphol 1965; 14(2):137-159. 130. Heller NH, Blecher SR. Reverse, hormone-dependent sex difference in molar tooth mass in pubertal mice. Arch Oral Biol 1982; 27(4):325-329. 131. Blecher SR. Anhydrosis and absence of sweat glands in mice hemizygous for the Tabby gene: Supportive evidence for the hypothesis of homology between Tabby and human anhydrotic (hypohydrotic) ectodermal dysplasia (Christ-Siemens-Touraine syndrome). J Invest Dermatol 1986; 87(6):720-722. 132. Cohen S. Isolation of a mouse submaxillary gland protein accelerating incisor eruption and eyelid opening in the newborn animal. J Biol Chem 1962; 237 1555-1562. 133. Blecher SR, Kapalanga J, Lalonde D. Induction of sweat glands by epidermal growth factor in murine X-linked anhydrotic ectodermal dysplasia. Nature 1990; 345:542-544. 134. Vargas GA, Fantino E, George-Nascimento C et al. Reduced epidermal growth factor receptor expression in hypohydrotic ectodermal dysplasia and Tabby mice. J Clin Invest 1996; 97(11):2426-2432. 135. Keen CL, Hurley LS. Developmental patterns of copper and zinc concentrations in mouse liver and brain: Evidence that the gene crinkled (cr) is associated with an abnormality in copper metabolism. J Inorg Biochem 1979; 11(3):269-277. 136. Keen CL, Hurley LS. Superoxide dismutase activity in the crinkled mutant mouse: Ameliorative effects of dietary copper supplementation. Proc Soc Exp Biol Med 1979; 162(1):152-156. 137. Hurley LS, Bell LT. Amelioration by copper supplementation of mutant gene effects in the crinkled mouse. Proc Soc Exp Biol Med 1975; 149(4):830-834. 138. Shawlot W, Siciliano MJ, Stallings RL et al. Insertional inactivation of the downless gene in a family of transgenic mice. Mol Biol Med 1989; 6(4):299-307. 139. Luetteke NC, Qiu TH, Peiffer RL et al. TGF alpha deficiency results in hair follicle and eye abnormalities in targeted and waved-1 mice. Cell 1993; 73(2):263-278. 140. Mann GB, Fowler KJ, Gabriel A et al. Mice with a null mutation of the TGF alpha gene have abnormal skin architecture, wavy hair, and curly whiskers and often develop corneal inflammation. Cell 1993; 2:249-261. 141. Luetteke NC, Phillips HK, Qiu TH et al. The mouse waved-2 phenotype results from a point mutation in the EGF receptor tyrosine kinase. Genes Dev 1994; 8(4):399-413. 142. Hu CC, Sakakura Y, Sasano Y et al. Endogenous epidermal growth factor regulates the timing and pattern of embryonic mouse molar tooth morphogenesis. Int J Dev Biol 1992; 36(4):505-516. 143. Canoun C, Ma C, Halpern D et al. Endogenous epidermal growth factor regulates limb development. J Surg Res 1993; 54(6):638-647. 144. Shum L, Sakakura Y, Bringas P Jr et al. EGF abrogation-induced fusilli-form dysmorphogenesis of Meckel’s cartilage during embryonic mouse mandibular morphogenesis in vitro. Development 1993; 118(3):903-917.

Ectodermal Dysplasia

37

145. Oosterwegel M, van de Wetering M, Timmerman J et al. Differential expression of the HMG box factors TCF-1 and Lef-1 during murine embryogenesis. Development 1993; 118:439-448. 146. Online Mendelian Inheritance in Man, OMIM (TM). Center for Medical Genetics, Johns Hopkins University (Baltimore, MD) and National Center for Biotechnology Information, National Library of Medicine (Bethesda, MD), 1996. World Wide Web URL: http:// www3.ncbi.nlm.nih.gov/omim/.

CHAPTER 3

Why Study Hair Follicles? A Personal Account Margaret Hardy Fallding

I

have had a long research career centered around hair follicles. I enjoyed it tremendously and so have some of my mentors and peers. Why do we do this? In this brief autobiographical commentary, I will make a personal reflection on my research career, spanning a period of 50 years. My intention is to share with you the joy I have in studying hair follicles, not to make a historic record on hair follicle research. My interest in hairs and hair follicles came about by accident. In my fourth year of the Honors B.Sc. degree program in Zoology at the University of Queensland, Professor Ernie Goddard sent me to Hayman Island in the Great Barrier Reef to collect all the different species of sponges (Porifera) that I could find, classify them, and write a thesis about them. This was a wonderful adventure in those days when the island was only a place for a handful of crazy fishermen living there in huts, and not the popular tourist paradise that it is now. By the time I had finished my thesis, I had learned that all the taxonomists who had worked on the sponges had drowned while collecting, become neurotic, manic depressive, or committed suicide! Professor Goddard then very wisely sent me off in the opposite direction—to Sydney to work with H.B. Carter, whom I will discuss more later on, at the F. D. McMaster laboratory in Sydney. This laboratory was part of the Commonwealth Scientific and Industrial Research Organization (CSIRO) and situated in the grounds of the University of Sydney— the best of all possible worlds! There I completed my M.Sc. thesis “Observations on the Mammalian Skin and Coat”, and then joined the CSIRO staff. When World War II ended, the CSIRO offered me a scholarship which enabled me to complete a Ph.D. in Zoology at the University of Cambridge and to visit many laboratories in the United Kingdom and in Europe. At the famous Strangeways Laboratory, the equally famous Honor B. Fell (later Dame Honor Fell) taught me her elegant techniques of whole organ culture with her own hands. I fell in love at once with the live, developing mouse hair follicle as I watched, through the microscope, how it developed from a tiny cluster of epidermal cells extending into the dermis. Then an even tinier cluster of dermal cells extended into the base of the lengthening follicle. When I was the first in the world to grow developing hair follicles in culture I told her at once. One hour later I walked into the tea room and was startled to find everyone in the Strangeways laboratory waiting to congratulate me! What a wonderful supportive environment to work in! I dreaded my thesis defense before two world-renowned External Examiners, with nobody else in the room, not even Miss Fell. It was almost a pleasure! After completing my Ph.D. thesis and on my way home to Australia through the USA, I visited the laboratory of Professor Charles Pomerat in the University of Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

40

Molecular Basis of Epithelial Appendage Morphogenesis

Texas at Galveston. I demonstrated my new techniques and learned some of theirs. Then I was back to the F. D. McMaster laboratory in Sydney, and began growing organ cultures of skin from fetal lambs. A few months after my marriage to Harold Fallding in 1954, I had to slow down on embryonic hair follicle culture because I was culturing an embryonic human, a much more difficult task for me! Then I had to stop my laboratory work in the early months of pregnancy and again one week before Marion was born. Three years later her sister Ruth came along, and Helen followed after another two years. I became a full-time mum. By the time I could get back to do part-time research it was 1963. We were in New Brunswick, New Jersey, where my husband was a Visiting Professor in Sociology at Rutgers University. I could not be at Rutgers because of existing anti-nepotism rules, so I had to commute to New York to the Columbia Medical School to join Dr. Margaret Murray and Dr. Richard Bunge, both tissue culture devotees. The trip was 1.5 hours each way. A few years later, my husband accepted a position as Founding Chair of Sociology at the University of Waterloo, Ontario. Again the prevailing anti-nepotism rules meant that I had to buy a car and drive to the University of Guelph, where I found a job as an Assistant Professor. I continued my research on hair follicles.1,2 Also, I continued the long distance driving until the end of 1996 when we moved to Guelph—a clear case of hair follicle addiction! My latest passion, shared with my present graduate student, Jennifer Robertson, is the use of immunohistochemistry to trace the biochemical changes in developing hair follicles. I have known many scientists with commitments similar to mine own to the wonders of the hair follicle. I have chosen several for particular mention in this brief autobiographical commentary, because I have collaborated with some of them and been greatly influenced by others. Many others whom I have not mentioned also have done important work in the field. I have enjoyed a good friendship with all, through our shared scientific interest. Dr. Harold Burnell Carter, B.V.Sc., never practiced veterinary medicine. He went straight from graduation to the F.D. McMaster Laboratory, CSIRO, because he was already in love with hair follicles. “H.B.”, as he came to be called, wanted to study the histology and histogenesis of wool follicles in sheep. He discovered that the first follicles in the sheep were arranged in rows of three, the trios, and usually produced the largest hairs. The later follicles clustered on one side of each trio, and produced the finer and shorter hairs. The trio follicles were named primary follicles, and the later, smaller ones were secondaries. Each primary follicle developed a small sweat gland and a pair of oil-secreting sebaceous glands. Each of the three glands opened into the neck region of its follicle. In addition, the primary follicles had a pair of smooth muscles which attached to the upper dermis in such a way that they could raise the sloping hair to an upright position on the skin surface. Some secondary follicles produced sebaceous glands, but none had sweat glands or smooth muscles. These features of individual hair follicles had been described earlier in some other species, but trio groups had not been described before. Carter studied large and small breeds of sheep from many parts of the world. He found a variety of densities (the numbers of follicles per unit area). The average number of follicles with fibers that began first (the primaries) on the whole animals of one breed tended to be fairly uniform, but the average numbers of secondary follicles between sheep varied. Thus the aim should be to breed for a higher ratio of secondary to primary follicles, rather than to aim for higher density. Some years later, the CSIRO, with funds from the wool industry, built a new laboratory on the outskirts of Sydney to accommodate the expanding research on wool including biochemistry, physiology, nutrition, genetics and breeding. Carter chose to work elsewhere in the CSIRO, and later on he moved to England and turned to writing about the histology of wool-growing, and doing work on skin and hair at the British

Why Study Hair Follicles?

41

Museum in London. H.B. continued with these activities with the same enthusiasm into his late eighties! What was it about skin and hair follicles that Carter found so fascinating? I think it was the beauty of the hair follicle itself and the layout of the patterns of follicle groups. Carter’s father was an artist, and H.B. had as sharp an eye as his father for beauty, whether it was a follicle group or a cluster of sailing boats on Sydney Harbor. Another person of great note and a close colleague who devoted her life to wool research was Miss Helen Newton Turner. She graduated from the University of Sydney, Australia, with a degree in Architecture, but since there were no jobs for architects during World War II, she worked as a secretary in the McMaster Laboratory! She typed research reports and scientific papers for publication. In order to understand what she was typing, she read a bit of Biology and of Statistics. Very soon this gifted person was the only official (parttime) Statistics Lecturer to the Veterinary School of the University of Sydney. She was also appointed as the Advisor on Statistics at the McMaster laboratory and told biologists, including me, which tests they should have been using! This woman whose only doctorate was an honorary one, later became Australia’s primary official Advisor to the Government on sheep and wool growth. Many countries invited her to advise them on crossbreeding and selection. She kept a diary during these travels and despite failing health, Helen kept on assembling and editing this diary for publication with the help of her niece, Loma Newton Priddle, until her death in her eighties in 1996.3 Dr. Francis William Dry recognized that sheep have distinct fiber types with particular shapes. For example, in the primary follicles, there were precurly tipped fibers, followed by fibers with sickle tips, then histiotrichs, all from the same follicles. Fibers with a coarse medulla were called halo hairs. F. W. Dry had his scientific education at Leeds and acknowledged his debt to his Zoology Professor Walter Garstang, and to Dr. R.S. Seton, the founder of the Department of Agriculture. His interest in wool was stimulated by Professor A.F. Barker of the Textile Industries Department at Leeds. Dry was a respectable English gentleman whose personality must have earned him the nickname of “Daddy Dry”. Many variations in fiber types from secondary follicles were seen by Dry and others, depending on the time of follicle formation. Each type of follicle had its own range of fiber types, and there were differences among the four seasons, among weather conditions and between individual sheep. Dry kept going with these studies from the nineteen-thirties to the fifties! In the sixties he was still at it, and he and others were still writing about follicles in the seventies. I treasure his 1975 book on the architecture of lambs’ coats.4 Another hair follicle devotee was Dr. R.I.C. Spearman, a British biologist who taught at University College Hospital Medical School in London, England. With a broad background in biology he wrote many papers about the outer skin layers of animals from Protozoa to the mammals. One of these was “The Integument”, published in the Symposia of the Cambridge University Press in 1973. In this book he reminds us that Aristotle, in the fourth century BC, described the details of male pattern baldness! Spearman explains how many advances in technology in the nineteenth century were first applied to skin and its appendages, in order to determine their structure and functions. Histological staining revealed the structural details of skin, glands, scales, feathers and hair follicles, and histochemical staining provided biochemical information. Later, phase contrast microscopy, radioisotopes and electron microscopy provided new revelations. Spearman held the view that it was the investigation of proteins and their structure in skin that opened up the whole field of molecular biology that we have today. The hair follicles that were collected as living structures, by noninvasive methods such as plucking, provided some of the earliest data of a biochemical nature in any tissue—the variety of keratin proteins, for example. I think Dr. Spearman

Molecular Basis of Epithelial Appendage Morphogenesis

42

stayed with the hair follicle for so long because it continued to yield useful information which was of value to biology in general.5 Dr. Philippe Sengel worked at the Université de Grenoble in the south of France, where he was Professeur de Biologie for many years. He quickly drew other scientists to his laboratory and to his subject of developmental biology. A superb writer and speaker in both French and English, he has contributed many important research papers on the development of skin, feathers and scales. He has also written several books which deal with the skin and skin appendages of mammals, birds and reptiles.6 Dr. Danielle Dhouailly joined the laboratory of Professeur Philippe Sengel after obtaining her first degree. There she became an expert in growing parts of chicken embryos on the chorioallantoic membranes of incubating eggs, and in recombining the separated epithelial and mesenchymal layers of embryonic skin on such a membrane. When I was on a sabbatical at Professeur Sengel’s laboratory in 1977, Danielle taught me these techniques. I taught her Honor Fell’s methods of in vivo organ-culturing and showed her the effects of addition of different types of retinoids on mouse skin organogenesis. In joint work Danielle and I found that injecting retinoids through the chorioallantoic membrane caused the scaleforming skin on the chicken embryos to be converted to feathers on the chicken feet!7 A major effect of this result was the conversion of Danielle to working with hair follicles (and retinoids!). She continues with organ cultures of skin and hair follicles and retinoids at Grenoble, and is now collaborating with molecular biologists to find out the molecular mechanisms of these findings. There are many other hair follicle addicts. Dr. George Rogers, for example, has spent all his working life studying the wonderful variety of keratin proteins, their genes and their different functions in different parts of the hair follicles and skin. He has also located these proteins at the ultrastructural level. He has been in demand as a speaker at conferences around the world, and has been instrumental in bringing hair and wool scientists together for conferences. Recently he was dreading his compulsory retirement from the Department of Biochemistry in the University of Adelaide in South Australia, but he was invited to continue his research in the Department of Animal Science as Professor Emeritus, with several assistants and good funding. In this discourse I have endeavored to show how the hair follicle has many ways of seducing scientists. It is tiny, beautiful, and yet it contains a lot of knowledge. Many challenges to understanding how follicles function still await our attention. The extensive knowledge of the mouse genome makes it a treasure trove for exploring skin and hair problems in mice, and this exploration may lead to treatments for other than animals, including Homo sapiens.

References 1. Hardy MH. Glandular metaplasia of hair follicles and other responses to vitamin A excess in cultures of rodent skin. J Embryol Exp Morphol 1968; 19:157-180. 2. Hardy MH. The secret life of the hair follicle. Trends Genet 1992; 8:55-61. 3. Priddle LN. ‘And Yonder Lies…’, Travels of a Remarkable Scientist. Glebe, Sydney, Australia: Fast Books, 1996. 4. Dry FW. The Architecture of Lambs’ Coats: A Speculative Study. Palmerston North, New Zealand: Massey University, 1975. 5. Spearman RIC. The Mammalian Epidermis and its Derivatives. Symp Zool Soc London 1964; 12:67-81. 6. Sengel P. Morphogenesis of Skin, Cambridge University Press, 1976. 7. Dhouailly D, Hardy MH, Sengel P. Formation of feathers on chick foot scales: a stagedependant morphogenetic response to retinoic acid. J Embryol Exp Morph 1980; 58:63-78.

Part II

Morphogenesis of Epithelial Appendages on the Body Surface

CHAPTER 4

Variations of Cutaneous Appendages: Regional Specification and Cross-Species Signals Danielle Dhouailly, Fabrice Prin, Benoit Kanzler and Jean P. Viallet

Introduction

I

n amniotes, the skin is well equipped with cutaneous appendages whose morphological type and distribution show specific regional variations. In birds, the integument can give rise to two types of appendages: scales and feathers. In chick, while most of the body is covered with feathers which are grouped in defined tracts or pterylae, the feet bear scales, and some areas, referred to as apteria, remain bare. In fact, two types of apteria must be distinguished: those that are entirely devoid of feathers (or scales), the midventral apterium and the comb; and those that show a few loosely distributed feathers, varying with each chicken, as for instance between the pectoral and alar tracts. The arrangement of feathers within each tract is constant and displays an open hexagonal pattern (see chapter 18). However, the outlines of each pteryla, as well as the number and shape of their constituent feathers, are region-dependent. Two major types of scales may be distinguished. Large scutate scales are arranged in two alternate longitudinal rows on either the anterior face of the tarsometatarsal or the proximal part of its posterior face, and in a single row on the upper face of each toe. Small roundish tubercular scales form a tight hexagonal pattern that covers the plantar face of the foot. Likewise, the hair follicles of mammals are not distributed, nor are they shaped at random. In mouse, the skin develops two major types of hair follicles. Pelage hair follicles of small diameter are arranged in triads and densely distributed, except on plantar regions, where they are mostly absent. Particularly, the foot pads are entirely devoid of hairs and made of a thick epidermis. In the facial region, the pelage hairs are intermingled with large sensory hair follicles or vibrissa which develop in rows around the mouth and the eyes. In this species, the tail epidermis shows a regular alternating pattern of parakeratotic scale regions and orthokeratotic interscale regions, with hairs erupted in a triad group at the distal end of each scale. Furthermore, in mammals another type of cutaneous appendage is well developed and diversified and forms the mammary, sebaceous and sweat glands. It should be noted that in mouse, in contrast to the human skin, the sweat glands are never associated with hair follicles but grouped on plantar foot pads. Thus, both in chick and mouse, the type and distribution of cutaneous appendages varies according to two axes: the antero-posterior (A-P) axis of the body, which is equivalent to the proximal-distal axis of the limb, and the dorso-ventral (D-V) axis. Morphologically, although the appearance of epidermal placodes precedes the visible patterning organization of dermal cells, the regional origin of the dermis controls their initiation, shape, and distribution, as shown by the results

Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

46

Molecular Basis of Epithelial Appendage Morphogenesis

of numerous heterotopic (different regions of skin), homospecific and heterospecific (different species) skin recombinations.1 Moreover, several experiments2,3 showed that the regional characteristics within different integumental regions are established very early during embryonic development, long before skin morphogenesis. In fact, recent experiments show that the different skin regions are specified within the establishment of the general body plan of the embryo, at the onset of organogenesis, and thus depend on the expression of both A-P and D-V signaling, which may interact. Transcription factors which specify the body plan, as well as secreted proteins which further allow the early steps of the continuous dialog between the two skin components, have not changed during amniote evolution and are thus understandable and correctly interpreted between a dermis and an epidermis belonging to two different classes.

The Formation of Embryonic Skin Varies According to the Different Body Regions In chick embryo, the skin forms at first in the back (future spinal pteryla) during the fifth day of incubation, and several days later in the head, the ventral face and the limbs. In mouse, the facial integument forms at 11 days of gestation, while the dorsal, limb and caudal skin morphogenesis begins respectively by 12.5, 14.5 and 15.5 days of gestation. Although only the avian model has been analyzed accurately, it is very likely that in this respect, avian and mammalian development do not differ significantly. In chick, the epidermis originates from the ectoderm but the nasofrontal skin area derives from the anterior part of the neural fold,4 whereas the origin of the dermis varies according to the different regions. The back dermis is formed by 5 days of incubation by the migration of somitic dermatomal cells,3 the ventral and limb dermis at 8.5 days by mesodermal cells which originate from the somatopleure. In fronto-nasal and branchial arc processes, the neural crest is the source of the connective tissue, as determined in chick5 and in mouse.6

The Predermal Cells Are Regionally Specifically Determined but Need a Systemic Nonspecific FGF-Like Epidermal Message to Acquire Their Dermal Inductive Potentialities The major part of our present knowledge of the early establishment of cutaneous patterns comes from studies of the chick embryo. Primary experiments7 have shown that the regional characteristics within different tracts and apteria are established very early during morphogenesis. Thus, at stage 21,8 the rotation of a block of superficial wing bud tissues (ectoderm plus mesoderm), corresponding to the future shoulder feather tract and to the region of the future elbow, results in shoulder tract deficiency and in a group of supernumerary feathers in the upper cubital apterium. Which of the two future skin components, ectoderm or mesoderm, is responsible for the alternative differentiation of an apterium or a pteryla? To answer this question, experiments have been performed consisting of heterotopic transplantations of blocks of ectoderm-free mesoderm.3 The results show that when a piece of lateral mesoderm from the prospective midventral apterium of a stage 16 embryo is implanted in the dorsal region, a patch of naked skin develops inside the dorsal tract. Furthermore, the specific characteristics of each region of the spinal pteryla are determined within the paraxial mesoderm.3 For example, when a strip of still unsegmented mesoderm from the thoracic region is transplanted into the posterior cervical region, the feather pattern that it elicits in the cervical epidermis is of the thoracic type. The dermis of the Scaleless mutant chick embryo, which is entirely devoid of scales and forms only a few feathers on the head and sacral regions, is regionally determined. Recent experiments from Song and coworkers9 showed that the mutant deficiency of the epidermis

Variations of Cutaneous Appendages

47

Fig. 4.1. Skin explants from four different body regions of Scaleless chick embryo, six days after treatment with FGF-2 loaded beads and graft onto chick chorioallantoic membrane. Induction of feathers (A) in E8 dorsal, of reticulate scales (B) in E9 plantar, of scutate scales (C) in E10 tarsometatarsal skin, whereas E10 midventral apterium skin remains glabrous (D).

can be recovered by bovine FGF-2 treatment. They obtained the formation of fused abnormally patterned short feathers on treated skin. In their experiments, the skin was cultured on a millipore filter placed onto the chorioallantoic membrane (CAM) of the chick embryo. By grafting the Scaleless skin directly onto the chick CAM before treating it with an appropriate dose of FGF-2, feathers arranged in an hexagonal pattern, reticulate scales, scutate scales and naked skin differentiated respectively onto dorsal (Fig. 4.1A), plantar (Fig. 4.1B), tarsometatarsal (Fig. 4.1C) and midventral apterium (Fig. 4.1D) skin explants.10 Thus, the wild-type FGF-producing epidermis may act as a systemic nonspecific homogeneous message, which allows the dermis to become organized according to its regional origin. Cutaneous regional specification appears thus to be determined long before skin formation. Nevertheless, skin regional morphogenesis requires further sequential and reciprocal interactions between the two skin components, the epidermis and the dermis.

Regional-Specific Dermal Messages Which Control the First Step of Cutaneous Appendage Formation Can Be Correctly Interpreted by an Epidermis from a Different Class The feather or hair rudiment comprises an epidermal circular placode which covers a dermal condensation. Although morphologically the appearance of epidermal placodes precedes the patterning organization of dermal cells,11,12 the regional origin of the dermis controls their initiation, shape, diameter and distribution.1 The dermo-epidermal interrelationship was investigated more than twenty years ago by recombining chick and mouse skin tissues either from feathered or scaled or haired areas or glabrous regions taken just before the appearance of appendage rudiments.1,13-15 From the chick embryo, dorsal, tarsometatarsal, plantar midventral and comb skin was used. From the mouse embryo, dorsal, upper lip and plantar skin was used. Skin heterospecific heterotopic recombinants were cultured for 6-8 days on the chick CAM. Recombinants of mouse plantar (non-hair-forming) epidermis with dorsal (featherforming) or tarsometatarsal (scale-forming) chick dermis differentiated typical stage 2-3 (as defined by Hardy in ref. 16) hair buds, whose development, however, did not proceed beyond this initial morphogenesis. Likewise, recombinants of midventral or comb (nonfeather-forming) chick epidermis with dorsal (pelage hair-forming) or upper lip (vibrissa hair-forming) mouse dermis formed arrested feathers, whose diameter and distribution (Fig. 4.2A,B) was determined by the regional origin of the mouse dermis. When the chick dorsal epidermis was associated with a mouse plantar dermis (from a 14.5 day embryo), the recombinant formed foot pads in their typical mouse pattern (Fig. 4.2C).17 The epidermis of these chimeric foot pads displays the numerous cell strata

48

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 4.2. Formation of cutaneous appendages in interclass dermal-epidermal recombinants between chick and mouse embryos. Formation of arrested feathers distributed in a typical hair vibrissae (A) or hair pelage (B) pattern in recombinants of 10 day chick comb epidermis and mouse 12.5 day upper lip (A) and 14.5 day dorsal (B) dermis. Formation of six foot pads (C) in their typical mouse pattern in a recombinant of 7 day chick dorsal epidermis and 14.5 day mouse plantar dermis. Note that the size and distribution pattern of the cutaneous appendages are dermis dependent. Histological sections (D, E) show the collaboration between chick and mouse cells. Formation of a feather bud (D) composed of a chick epidermal sheath overlying a mouse dermal condensation when a 6 day chick dorsal epidermis is associated with a 14.5 day mouse dorsal dermis. Formation of a hair bud (E) developed in contact with a chick dermal condensation when a 12.5 day mouse dorsal epidermis is associated with a 7 day chick dorsal dermis.

which are characteristics of the mouse foot pads. Moreover, those cells synthesize scalespecific β keratin polypeptides. Thus it can be advanced that the chick dorsal epidermis, which normally would have formed feathers and their specific β keratins, was induced to form pads which are morphologically related to scutate scales. This event would then dictate the synthesis of the appropriate keratin polypeptides. Likewise, contrary to an ancient hypothesis,18 the chick tarsometatarsal dermal cells are invested with proper scale-forming properties before 12 days of incubation, as demonstrated by the association of tarsometatarsal dermis with an apteric epidermis.19 In contrast, the heterotopic homospecific recombination of the chick dorsal epidermis with a midventral apterium dermis did not form cutaneous appendages, but an apterous skin.20 As the formation of a specific appendage tract or an apterium is dependent on the regional origin of the mesoderm in heterotopic transplantation experiments, it seems reasonable to presume that it results from autonomous dermal cell capacities, which are dependent on a permissive FGF-2-like epidermal message (see previous paragraph). Within each tract, what are the mechanisms governing the subdivision of the dermis? A partial answer to this question is discussed in chapter 18. What is the nature of the messages which are transmitted between the epidermis and the dermis at the first step of cuta-

Variations of Cutaneous Appendages

49

neous appendage morphogenesis, when the two skin components are able to cooperate perfectly even if they belong to two different classes of vertebrates (Fig. 4.2D,E)?13 The secreted molecules which allow the condensation of the dermal cells and the growth of the placode into a bud are well conserved between the different classes of vertebrates and belong to the hedgehog and TGF families, as shown by Chuong and coworkers.21-23 This question is examined in more detail in chapter 13. It should be noted that the development of cutaneous appendages obtained in inter-class recombinants does not proceed beyond the bud stage, suggesting that a second step of dermal induction is necessary.1 Another argument in favor of the necessity of a two step induction is the fact that dermal cells from glabrous regions which are unable to exert the first initiating induction are in contrast able to support the continuation of appendage morphogenesis, as shown by the results of heteroclass (different classes of vertebrates) recombinants involving a mixed dermis with a few dermal cells from a glabrous region belonging to the same class as the epidermis.24 For example, small arrested feather filaments are grouped together with a few normal feather filaments with well organized barb ridges when a dorsal quail epidermis is associated with a chimeric bispecific dermis composed of 90% dorsal mouse epidermis and 10% of midventral apterium chick dermis.24

The Epidermis Does Not Possess Equivalent Competence All Over the Body Not only the dermis, as previously thought,1,25,26 but also the epidermis plays an important role as a regional agent during skin morphogenesis. Heterotopic recombinants of ectoderm and mesoderm of wing and leg buds of 4 day chick embryos result in the development of a limb, the regional quality of which is in accordance with the regional origin of the mesoderm. The leg ectoderm differentiates into normal feathers when it is recombined with a wing mesoderm. However, the reverse combination of wing ectoderm and leg mesoderm leads to the formation of feet covered with feathers only or with feathered scales.27 From these results, it is clear that the ectoderm of the wing bud and that of the leg bud are not equivalent with respect to their cutaneous appendage competence. Likewise, when 7 day dorsal epidermis was associated with a 10 day plantar dermis, it developed numerous and long feather filaments, whereas the reverse association of a 10 day plantar epidermis with a dorsal dermis leads to the formation of arrested feathers.28 Thus the plantar dermis possesses quite similar feather inductive abilities as the dorsal dermis, but the feather morphogenesis is in some way inhibited in the plantar epidermis. This is confirmed by the fact that the wild-type or Scaleless plantar skin cannot be induced by FGF-2 treatment to form feathers.10 In contrast, the avian extraembryonic somatopleure (the ectoderm plus its underlying mesoderm) is imprinted with feather-forming capacities.29 In mouse, by 12 days of gestation, the dorsal and upper lip epidermis also have different potentialities. Although upper lip and dorsal dermis are able to induce respectively the differentiation of hair vibrissa and hair pelage follicles in a dorsal or an upper lip epidermis,15 when the same types of recombinants are treated with retinoic acid, only those which involve an upper lip epidermis form glandular structures instead of hair vibrissae.30 The glandular metaplasia of the upper lip skin, first studied by Hardy,16,31 is thus due to a particular competence of the facial epidermis and is not linked to a particular hair morphology. The effect of retinoic acid, as shown by Blanchet and coworkers32 is principally monitored by its nuclear receptor RAR γ, which is the main receptor to be expressed in the epidermis of different species.33-36 By using a specific agonist, which unlike the retinoic acid molecule, is neither light sensitive nor rapidly metabolized, even 100% of the vibrissae are transformed into glands.32 What is the molecular basis of the different epidermal potentialities, both in chick and mouse embryos? Those differences, which occur at early stages of

50

Molecular Basis of Epithelial Appendage Morphogenesis

embryogenesis, can either be autonomous and thus correspond to a variability in competence, or result from early mesenchymal inductive stimuli.

Molecular Specification Along the Anteroposterior and Dorsoventral Axes Act Together Toward Skin Regionalization Results in the early nineties have demonstrated that distinct homeobox gene families are responsible for the regional diversity of the brain37,38 and of the axial structures39-41 or limb.42 What is the distribution of those genes expressed in the two skin components and do they play a role in skin regionalization? Vertebrate homeobox genes, which are homologous to the first identified genes of the Drosophila Antennapedia and bithorax complexes are organized into four paralog gene clusters, HoxA, B, C and D, located on different chromosomes (for reviews see refs. 40 and 43). Within each cluster, genes most similar in sequence to a particular Drosophila gene, or paralog genes, occupy the same relative positions within their respective cluster. In addition, the rostro-caudal boundary of one given Hox expression domain correlates with the position of the gene within its cluster, a property termed colinearity.44,45 The same Hox sets have been shown to regulate the developmental processes within antero-posterior and proximo-distal specification in patterning of the vertebrae and limb segments (among others, refs. 39 and 42). For skin, first results showed that the Hoxc6 and Hoxd4 homeoproteins are differentially expressed during dorsal chick skin formation.21,22 It is well established that during embryonic vertebrate development, the expression of the Hox genes does not occur beyond the second rhombomere (among others, see ref. 46). Two other homeobox-containing gene classes were primarily isolated in mouse,37,47 which have restrictive expression areas in the developing brain: the Otx1 and Otx2 genes, related to the orthodenticle Drosophila gene; and the Emx1 and Emx2 genes, related to the empty spiracle Drosophila gene. We showed48 that in mouse skin, the Otx and the Hox genes identify the facial and body territories respectively. Indeed, the Otx2 and Otx1 transcripts appear first during mouse skin formation and are restricted respectively to the nasal and facial epithelia, whereas Hoxc8 (Fig. 4.3B), Hoxd9, Hoxd11 (Fig. 4.3D) and Hoxd13 (Fig. 4.3F) transcripts appear later and successively and identify respectively the thoracic, lumbar, caudal and plantar skin. Among the different Hox genes studied, the transcripts are mostly limited to the epidermal component of the skin, and are not expressed once cutaneous appendage morphogenesis is accomplished, except in the epidermal basal layer. However, Hoxc8 is expressed both in the epidermis and the upper dermis, while Hoxd13 expression is limited to the dermis. Similar studies have been performed in chick embryo.29 The developmental expression pattern of the Hoxc8 and Hoxd13 genes suggest that these homeoproteins play a role in the specification of dorsal and plantar skin morphogenesis, as they are expressed at the first stages of feather and reticulate scale formation, in, respectively, the dermis and the epidermis (Hoxc8) (Fig. 4.4A) and only the dermis (Hoxd13) (Fig. 4.4C). The Hoxd11 gene is not expressed in the skin of the short tail of the chicken embryo (Fig. 4.3C), while its transcripts mark the mouse embryo caudal skin (Fig. 4.3D). Hoxd13 expression characterizes, in both chick and mouse, the ventral skin of both wing and leg autopodes when skin differentiation takes place, particularly the plantar dermis. The only difference is that in the mouse, the Hoxd13 gene is expressed in a slightly more distal skin region, namely in the anterior and plantar dermis from the digits (unpublished data from Viallet). Thus the restricted expression pattern of Hox genes which occurs during the first stages of skin morphogenesis appears to be very similar in both mouse and chick embryos. Given their distinct expression pattern in skin, the Hoxc8 and Hoxd13 genes could be part of different homeoprotein sets, providing positional information specifying the thoracic and autopodial skin territories respectively. The results of heterotopic recombinants29 show that

Variations of Cutaneous Appendages

Fig. 4.3. Differential expression of Hoxc-8, Hoxd-11 and Hoxd-13 in the skin of chick (A, C, E) and mouse (B, D, F) embryos. Distribution of the Hoxc8 transcripts at the onset of dorsal skin morphogenesis in sagittal sections of (A) 7.5 day chick and (B) 14.5 day mouse embryos (rostral at the left). The transcripts display a similar anterior expression boundary (arrowhead) in thorax, both in the skin (s), the vertebrae (vt) and the neural tube (nt). Distribution of the Hoxd11 transcripts in sagittal sections of the caudal region of a 7.5 day chick embryo (C) and a 16.5 day mouse embryo (D). In both species, the transcripts are present in the neural tube and the vertebrae. The mouse caudal skin is labeled with an anterior limit (arrowhead) posterior to the expression in vertebrae and neural tube, which can explain the absence of labeling of the chick caudal skin, as there are only four caudal vertebrae in this species. Distribution of Hoxd13 transcripts in longitudinal sections of the foot of a 10.5 day chick (E) and a 17.5 day mouse (F) embryo. The transcripts are present in the perichondrial mesenchyme and the plantar skin (ps).

51

52

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 4.4. Hoxc8, Hoxd11 and Hoxd13 are only expressed in the early stages of cutaneous appendage morphogenesis. For example, the Hoxc8 (A, B) transcripts are present in the dorsal feather primordia (A), both in the epidermis (e) and the dermal condensation (dc), while they are not detectable in the feather filament (f) (B) (the labeling indicated by the arrowhead, which is confined to the base of the feather filament, is an artefact). The Hoxd13 transcripts are present at the onset of plantar reticula morphogenesis (C) and restricted in the skin to the dermal cells (d), and not detectable when the reticula are formed (D). Likewise, The Hoxd11 transcripts are present in all epidermal cells of the caudal hair buds (hb) (E), as well as in the basal cells (bc) of the epidermis, and are restricted to the basal cells of both the outer root sheath (ors) and of the epidermis when the hair follicle (hf) is formed (F).

the plantar epidermis acquires a restricted ability to differentiate only reticulate scales, which coincides with the expression of Hoxd13 in the underlying dermis, while the dorsal epidermis, which expresses Hoxc8, is endowed with feather forming ability, as shown by its ability to interpret inducing clues originating either from a dorsal or a plantar dermis to go through the feather differentiation program. At the time of chick skin recombination, 7 days for the dorsal, and 11 days for the plantar tissues, the Hox genes have already played their role in specifying epidermal abilities. This probably even occurs earlier, when the global pattern of Hox gene expression within the body is established in early embryos. Nevertheless, the difference between the ventral skin of the wing, which is feathered, and the plantar skin, i.e., the formation of reticulate scales, must involve a further set of information which prevents the plantar epidermis from developing into feathers.10

Variations of Cutaneous Appendages

53

As it is well known that retinoic acid (RA) is able both to modulate homeobox gene expression49,50 and to promote the ectopic formation of feathers on chick feet,51 we repeated this RA treatment in order to analyze the possible correlative changes in Hoxd13 expression. Retinoic acid treatment at 10 or 11 days of incubation resulted in an inhibition of Hoxd13 expression in the plantar dermis. Embryos treated on day 11 developed feather filaments on the reticulate scales. In contrast, in embryos treated on day 10, the Hoxd13 expression reappeared in 24 h, followed by a normal reticulate scale morphogenesis. A possible explanation is that retinoic acid acts by inhibiting the posterior paralogs and respecifying the plantar cells to a more proximal positional identity, thus allowing the formation of feathers. It cannot be excluded that RA may also act by enhancing the expression of more anterior paralogs. It should be noted that these RA effects on embryonic skin are consistent with the known effects of RA on Hox gene expression in teratocarcinoma cells in vitro49,50 during vertebrae differentiation39 and limb morphogenesis:52 the expression of Hox genes belonging to the median paralogs 4 to 8 is generally not modified by RA, whereas the 5' clustered genes are downregulated. Nevertheless, the specification of skin regionalization must require, in addition to the proximo-distal or antero-posterior clues provided by the Hox genes, still unknown information involved in the characterization of the hind limb versus the wing, as well as some information to specify the position on the dorso-ventral axis. Recent studies have identified a number of molecules involved in the dorso-ventral signaling and there is increasing evidence that the signaling systems interact with one another.53,54 Patterning along the D-V axis is controlled by signals from the ectoderm. Thus, rotation of limb bud ectoderm induces a corresponding reversal in skeletal, muscle and skin pattern.55 Signaling from dorsal ectoderm is mediated by a member of the Wnt family of secreted proteins, Wnt7a.56 Loss of Wnt7a function in homozygotic mutant mice results in the transformation of the dorsal skin: intermingled with the pelage hairs which cover the dorsal surface of the paw, ectopic plantar foot pads formed. Furthermore, Wnt7a appears to mediate dorsalization of the mesenchyme through induction of the LIM homeobox containing gene, Lmx1.53 Surprisingly, ectopic Lmx1 expression causes changes of the foot integument in chick embryo from reticulate scales to feathers, and not, as expected, from reticulate to scutate scales. Mutational analysis in mice has shown that the homeobox containing the gene Engrailed 1 is involved in specifying the ventral side. Loss of Engrailed 1 function results in dorsalization of the foot pads, but not in a thinning of the hair pelage of the dorsal surface of the mouse paw.54

Conclusion Thus it is no longer surprising that epidermal and dermal tissues belonging to two different classes of vertebrates such as the chick and the mouse can collaborate during the early steps of cutaneous appendage formation. Indeed, the specification of their regional characteristics are based on common mechanisms which involve the same transcription factors and secreted proteins. The different areas of vertebrate integument are specified together with the general body plan in the early embryo. During the second step of cutaneous appendage morphogenesis, e.g., the construction of a feather or a hair, discrepancies appear among the different vertebrate classes. Probably the same adhesive molecules and growth factors are still used, but their pattern of expression must depend on still unknown, and different, mechanisms.

References 1. Dhouailly D. Dermo-epidermal interactions during morphogenesis of cutaneous appendages in amniotes. Front Matrix Biol 1977a; 4:86-121.

54

Molecular Basis of Epithelial Appendage Morphogenesis

2. Saunders JW, Gasseling MT, Cairns JM. The differentiation of prospective thigh mesoderm grafted beneath the apical ectodermal ridge of the wing bud in the chick embryo. Dev Biol 1959; 1:281-301. 3. Mauger A. Rôle du mésoderme somitique dans le développement du plumage dorsal chez l’embryon de poulet. II Régionalisation du mésoderme plumigène. J Embryol Exp Morphol 1972; 28:343-366. 4. Couly G, Le Douarin N. The fate map of the cephalic neural primordium at the presomitic to the 3-somite stage in the avian embryo. Development 1988; 103s:101-113. 5. LeLièvre CS, LeDouarin N. Mesenchymal derivatives of the neural crest: Analysis of chimeric quail and chick embryos. J Embryol Exp Morphol 1975; 34:125-154. 6. Osumi-Yamashita N, Ninomiya Y, Doi H et al. The contribution of both forebrain and midbrain crest cells to the mesenchyme in the frontonasal mass of mouse embryo. Dev Biol 1994; 164:409-419. 7. Saunders JW, Gasseling MT. The origin of pattern and feather germ tract specificity. J Exp Zool 1957; 135:503-528. 8. Hamburger V, Hamilton HL. A series of normal stages in the development of the chick embryo. J Morph 1951; 88:49-92. 9. Song H, Wang Y, Goetinck PF. Fibroblast growth factor 2 can replace ectodermal signaling for feather development. Proc Natl Acad Sci USA 1996; 93:10246-10249. 10. Prin F, Olivera-Martinez I, Viallet JP et al. Fibroblast growth factor 2 allows the regional organization of the chick dermis. J Int Dev Biol In Press. 11. Sengel P, Rusaouën M. Aspects histologiques de la différenciation précoce des ébauches plumaires chez le poulet. C R Acad Sci Paris 1968; 266, 795-797. 12. Dhouailly D. Specification of feather and scale patterns. In “Pattern formation“, Malincinski GM, Bryant SV, Eds. New York: Macmillan Pub Co 1984: 581-601. 13. Dhouailly D. Dermo-epidermal interactions between birds and mammals: Differentiation of cutaneous appendages. J Embryol Exp Morphol 1973; 30:389-400. 14. Dhouailly D. Formation of cutaneous appendages in dermo-epidermal interactions between reptiles, birds and mammals. Roux Arch Dev Biol 1975; 177:323-340. 15. Dhouailly D. Regional specification of cutaneous appendages in mammals. Wilhelm Roux’ Archives 1977b; 181:3-10. 16. Hardy MH. Glandular metaplasia of hair follicles and other responses to vitamin A excess in cultures of rodent skin. J Embryol Exp Morphol 1968; 19:157-180. 17. Dhouailly D, Rogers GE, Sengel P. The specification of feather and scale protein synthesis in dermal-epidermal recombinations. Dev Biol 1978; 65:58-68. 18. Rawles ME. Tissue interactions in scale and feather development as studied in dermalepidermal recombinations. J Embryol Exp Morphol Exp 1963; 11:765-789. 19. Cadi R, Dhouailly D, Sengel P. Use of retinoic acid for the analysis of dermal-epidermal interactions in the tarsometatarsal skin of the chick embryo. Dev Biol 1983; 100:489-495. 20. Sengel P, Dhouailly D, Kieny M. Aptitude des constituants cutanés de l’aptérie médioventrale du poulet à former des plumes. Dev Biol 1969; 19:436-446. 21. Chuong C-M, Olivier G, Ting SA et al. Gradients of homeoproteins in developing feather buds. Development 1990; 110:1021-1030. 22. Chuong C-M, Widelitz RB, Ting-Xin J. Adhesion molecules and homeoproteins in the phenotypic determination of skin appendages. J Inv Derm 1993; 101:10S-14S. 23. Chuong C-M, Widelitz RB, Ting-Berreth S et al. Early events during avian skin appendage regeneration: dependance on epithelial-mesenchymal interaction and order of molecular reappearance. J Inv Derm 1996; 107:639-646. 24. Dhouailly D, Sengel P. Propriétés phanérogènes des cellules dermiques de peau glabre d’oiseau ou de mammifère. C R Acad Sci Paris 1975; D 281:1007-1010. 25. Sengel P. Morphogenesis of skin. In Developmental and Cell biology series. Abercrombie M, Newth DR, Torrey JG, Ed.; Cambridge: University Press 1976:1-277. 26. Sawyer RH. The role of epithelial-mesenchymal interactions in regulating gene expression during avian scale morphogenesis. In “Epithelial-mesenchymal Interactions in Development”. Sawyer RH, Fallon, JF Eds. New York: Praeger Press 1983:115-146.

Variations of Cutaneous Appendages

55

27. Sengel P, Pautou MP. Experimental conditions in which feather morphogenesis predominates over scale morphogenesis. Nature 1969; 222:693-694. 28. Kanzler B, Prin F, Thélu J et al. cHoxc-8 and cHoxd-13 expression in embryonic chick skin and cutaneous appendages specification. Dev Dyn 1997; 210:274-487. 29. Dhouailly D. Feather-forming capacities of the avian extra-embryonic somatopleure. J Embryol Exp Morphol 1978; 43:279-287. 30. Viallet JP, Dhouailly D. Retinoic acid and mouse skin morphogenesis: II. Role of epidermal competence in hair glandular metaplasia. Dev Biol 1994a; 166:277-288. 31. Hardy MH. Vitamin A and the epithelial-mesenchymal interactions in skin differentiation. In “Epithelial-mesenchymal Interactions in Development”. Sawyer RH, Fallon JF Eds. New York: Praeger Press 1983: 163-188. 32. Blanchet S, Chevalier G, Kastner P et al. Differential roles of retinoid receptors in retinoid-induced mouse upper-lip skin glandular metaplasia. J Inv Derm 1998; (in press). 33. Petkovich M, Branb NJ, Krust A et al. A human retinoic acid receptor which belongs to the family of nuclear receptors. Nature 1987; 330:444-450. 34. Viallet JP, Dhouailly D. Retinoic acid and mouse skin morphogenesis. I. Expression pattern of retinoic acid receptor genes during hair vibrissa follicle, plantar and nasal glands development. J Inv Derm 1994b; 103:116-121. 35. Michaille JJ, Blanchet S, Kanzler B et al. Characterization of cDNAs encoding the chick retinoic acid receptor γ2 and preferential distribution of retinoic acid receptor γ transcripts during chick skin development. Dev Dyn 1994; 201:334-334. 36. Michaille JJ, Kanzler B, Blanchet S et al. Characterisation of cDNAs encoding two chick retinoic acid receptor isoforms and distribution of retinoic acid receptor α, β and γ transcripts during chick skin development. Int J Dev Biol 1995; 39:587-596. 37. Simeone A, Acampadora D, Gulisano M et al. Nested expression domains of four homeobox genes in developing rostral brain. Nature 1992a; 358:687-690. 38. Boncinelli E, Gulisano M, Broccoli V. Emx and Otx homeobox genes in the developing mouse brain. J Neurobiol 1993; 24:1356-1366. 39. Kessel M, Gruss P. Homeotic transformations of murine vertebrae and concomitant alteration of Hox codes induced by retinoic acid. Cell 1991; 67:89-104. 40. McGinnis W, Krumlauf R. Homeobox genes and axial patterning. Cell 1992; 68:283-302. 41. Le Mouellic H, Lallemand Y, Brûlet P. Homeosis in the mouse induced by a null mutation in the Hox-3.1 gene. Cell 1992; 69:251-264. 42. Duboule D. The vertebrate limb: a model system to study the Hox/HOM gene network during development and evolution. Bio Assays 1992; 14:375-383. 43. Krumlauf R. Hox genes in vertebrate development. Cell 1994; 78:191-201. 44. Duboule D, Dolle P. The structural and functional organization of the murine HOX gene family resembles that of Drosophila homeotic genes. EMBO J 1989; 8:1497-1505. 45. Duboule D, Morata G. Colinearity and functional hierarchy among genes of the homeotic complexes.Trends Genet 1994; 10:358-364. 46. Krumlauf R. Hox genes and pattern formation in the branchial region of the vertebrate head. Trends Genet 1993; 9:106-112. 47. Simeone A, Gulisano M, Acampadora D et al. Two vertebrate homeobox genes related to the Drosophila empty spiracles gene are expressed in the embryonic cerebral cortex. Embo J 1992b; 11:2541-2550. 48. Kanzler B, Viallet JP, Le Mouellic H et al. Differential expression of two different homeobox gene families during mouse tegument morphogenesis. Int J Dev Biol 1994; 38:633-640. 49. Simeone A, Acampora D, Nigro V et al. Differential regulation by retinoic acid of the homeobox genes of the four HOX loci in human embryonal carcinoma cells. Mech Dev 1991; 33:215-227. 50. Boncinelli E, Simeone A, Acampora D et al. HOX gene activation by retinoic acid. TIG 1991; 7:329-334. 51. Dhouailly D. Hardy MH, Sengel P. Formation of feathers on chick foot scales: a stagedependant morphogenetic response to retinoic acid. J Embryol Exp Morph 1980; 58:63-78.

56

Molecular Basis of Epithelial Appendage Morphogenesis

52. Hayamizu TF, Bryant SV Reciprocal changes in Hoxd-13 and RAR-β2 expression in response to retinoic acid in chick limb buds. Dev Biol 1994; 166:123-132. 53. Vogel A, Rodriguez C, Warnken W et al. Dorsal cell fate specified by chick Lmx1 during vertebrate limb development. Nature 1995; 378:716-720. 54. Loomis CA, Harris E., Michaud J et al. The mouse Engrailed-1 gene and ventral limb patterning. Nature 1996; 382:360-363. 55. MacCabe JA, Errick J, Saunders JW. Ectodermal control of the dorsoventral axis in the leg bud of the chick embryo. Dev Biol 1974; 39:69-82. 56. Parr BA, McMahon AP. Dorsalizing signal Wnt-7a required for normal polarity of D-V and A-P axes of mouse limb. Nature 1995; 374:350-353.

CHAPTER 5

Feather Morphogenesis: A Model of the Formation of Epithelial Appendages Cheng-Ming Chuong and Randall B. Widelitz

Introduction

A

mong the skin appendages, feathers certainly have the most elaborate structures and are richly patterned. Feathers distinguish the bird from other vertebrate classes and provide the birds with a unique niche, the sky, in which to thrive and evolve. Feathers also serve other crucial functions such as thermal insulation, communication, the ability to swim, water-repellence, support, camouflage, etc. The feather indeed becomes a very important part of the life of a bird. There are different types of feathers (Fig. 5.1).1 The major ones are contour feathers that are made of a central stiff branch, the rachis, with barbs branching out from the rachis, and barbules branching from the barbs. The distal edge of the barbule has small hook projections (barbicels) that can interlock with the proximal edge of the next barbule. This velcro-like mechanism produces a strong, flexible, light-weighted planar structure named the vane. A flight feather from the pigeon has 600 pairs of barbs and each barb has 500 barbules. On the average, a feather has approximately 2400 of these tiny zippers.2 The other major type is down feathers that are soft and fluffy. They tangle with each other and trap air to form an excellent thermal insulator. Filoplumes are hair-like feathers that can monitor the position of other feathers and are distributed near mechanically active feathers. Bristles are made of a long rachis with very few barbs. They are found around the mouth of some owls, and assume a whisker-like function. There are also minor types and numerous modifications of the feathers. The range of feather designs in the Aves class can dazzle fashion designers. The feather is a major organ in the body of a bird. An average swan can have up to 25,000 feathers, and a song bird typically has around 2000-4000 feathers. The size of feathers ranges from 1 mm to longer than 1 m. Although individual feathers are light, their combined weight contributes tremendously to the mass of a bird. Using a 4 kg bald eagle as an example, the total skeleton weighs 272 gm, or 7% of the body mass, while the total mass of its feathers is 700 gm or 17% of the body mass. Each feather is rooted in the dermal region with a follicular structure (Fig. 5.2) which is fixed to the dermis by dense connective tissues and muscles.3 Feather movement is regulated through connections between the follicles and an intricate neuromuscular network that coordinates the movement of the approximately 20,000 feathers on the body. At the base of the follicle is the dermal papilla. The dermal papilla is surrounded by the papillary ectoderm that generates new epithelial cells Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

58

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 5.1. Schematic drawing to show the variety of feather morphology. Basic morphological elements of feathers are shown. The proportions of the lengths of the rachis, barbs and barbule, and the angles between barbs and rachis, θ, can vary, giving feathers different shapes and specialized functions. (A) is a typical bilateral symmetric contour feather. With interlocking barbules (shown in the right upper vane), the feather becomes a plane and can be strong while maintaining light weight. (B) is an asymmetric feather seen in tail feathers and flight feathers. Both the length of barbs and the angle with the rachis, θ, are different in the left and right vanes. (C) is fan shaped, frequently seen in trunk and head. (D) is a fluffy down feather, good for keeping warmth. (E) has very short barbs and barbules and is hair-like, as seen in penguins. (F) is the basic structure of filoplumes or bristles, which can have some tactile functions. Some feathers can have a mixture of two types, such as a tightly interlocking distal vane and a downy proximal base. The space between barbs is originally occupied by the marginal plate (mp), and the space between barbules is originally occupied by the axial plate (ap). Please refer to Fig. 5.2.

upon molting or regeneration. Above the dermal papilla is the collar epithelium that contains actively proliferating cells. This region is equivalent to the hair matrix and probably contains transient amplifying cells.4 The epithelial cylinder cells advancing from the upper collar region generate obliquely-arranged alternating compartments that keratinize or apoptose to form feather branches and the inter-branch space. The feather pulp occupies the transient space originally enclosed by the epithelial cylinder. It contains loose connective tissues, blood vessels and nerves, and disappears when feathers mature. The distinct shapes and sizes of the feather make it a spectacular organ for the study of morphogenesis and a model for the formation of epithelial appendages. Feathers do not function alone; rather they are organized in groups to effectively carry out their function. Regions of grouped feathers are named tracts, or pterylae. Feather morphology is specified differently in different feather tracts (Fig. 5.3A-C).5 On each tract, feathers grow in a coordinated order with characteristic spacing and orientation (Fig. 5.3D). For example, in the spinal tract, developing feather buds are arranged in a regular hexagonal pattern. Feather arrangement has been a favored model for the study of pattern formation (Fig. 5.4). How does a presumably flat piece of skin generate the diverse structures found in different feather tracts, and in different species (Fig. 5.1)? Differences between species can be attributed to genetic differences generated in evolution (see chapter 19). However, in a single bird the differences among the variety of feathers and scales on the integument are still

Feather Morphogenesis

59

Fig. 5.2. Three dimension view of the topological transformation of a feather from a two dimension flat sheet to a three level branched structure. (A-E), Topological transformation from a two dimensional plane to a branched feather filament. (F), A three dimension view showing how the surface of the feather filament is “sculptured” into a highly branched structure. (G) shows the base of the follicle. For simplification, barbules are not shown. cl, collar; dp; dermal papilla; fs, follicular sheath; p, pulp; pe, papilla ectoderm. Panel (A-E) is modified from Chuong and Edelman, 1985;19 Panel (F) is modified from Lucas AM and Stettenheim 1972.3

60

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 5.3. Overview of developing feather buds. (A), Stage 29. (B) and (C), stage 36, dorsal and ventral view. The distinct arrangement and shape of tracts constitute the macropattern. The distinct arrangement of feather buds within each tract constitute the micropattern. Note that the tract begins as a homogeneous tract field (A) from which periodically arranged feather primordia emerge sequentially from the primary row (B), (also see Fig. 5.4). AT, alar tract; AbT, abdominal tract; CT, cervical tract; CaT, caudal tract; FT, femoral tract; HT, humeral tract; ST, spinal tract; StT, sternal tract. (D), A piece of dorsal skin from a stage 37 embryo hybridized to an Shh antisense probe.34 This is used to show feather primordia of different developmental stages. From the midline (diagonal line from the left lower corner to the right upper corner) to both lateral edges are long feather buds, short feather buds, and feather placode. Shh are on the tip of the placode and distal epithelium of the short bud. In the long bud, Shh is also expressed on the marginal plate epithelium, forming a striking stripe pattern. See color insert.

Feather Morphogenesis

Fig. 5.4. Schematic drawing of the development of feather tracts and feather buds. (AC), Development of feather tracts. (A), dorsal body surface. (B), The appearance of the primary row and tract field in the spinal tract (ST), two humoral tracts (HT) and two femoral tracts (FT). (C), The emergence of individual feather primordium. (D), Development of feather buds. There are different stages in the development: 1. Competent cells in the tract field start the initiation process of feather primordia; 2. Setting up the boundary of dermal condensation; 3. Asymmetry formation, or setting up the anterior posterior (A-P) axis; 4. Elongation, or the formation of proximal-distal (P-D) axis; 5. Morphogenesis of feather filament branches, namely barbs and barbules.

61

62

Molecular Basis of Epithelial Appendage Morphogenesis

complex. These different structures are presumably derived from the same genetic information, regulated under different epigenetic control. Classical embryologists have been fascinated by these exquisite morphologies, and have tried to understand the factors that regulate diverse formative processes using experimental embryology and transplantation experiments (please see chapter 4). In many cases, they have successfully localized the determination factors to the mesenchyme.6-8 With the advent of molecular biology, today we are trying to find the molecular basis for these phenomena. It is the goal of this chapter to present an overview of feather morphogenesis, to review what we know in each phase of feather development and to point out areas for future study. We hope that some of these concepts may enhance our understanding of the general principles of epithelial appendage construction.

Overview of Feather Morphogenesis Feather formation is basically a topological transformation from a two dimensional plane to an elaborate three-dimensional branched skin appendage (Figs. 5.2-5.6). This developmental progression can be divided into different phases (Fig. 5.4).9 First, a field competent to form feathers is established and this field becomes the future feather tract. Second, the site of an individual feather primordium within a feather field is set. Third, the orientation of the feather primordium is determined. Fourth, cells, the raw material for building a feather, are generated by cell proliferation. Finally, the filament is "carved" into branched structures. During these five phases of feather morphogenesis, different cellular and molecular morphogenetic events occur. To analyze these processes, we define the stages based on morphological characteristics and the state of determination (Fig. 5.7). We have also begun to place some known molecular markers along this staging system. Some of these staging criteria have been described earlier,7,10 but are reviewed here with modifications. We will first present a glossary of terms to be used in this chapter. Feather primordia is used loosely to include all stages of developing feathers. Feather rudiment is used to indicate the earliest events in feather morphogenesis. It includes initial events that are undetectable with current techniques, such as those beginning before the placode is visible. Feather placode is characterized by the elongation of epithelial cells in the domain where a feather primordium will form. Dermal condensation is the accumulation of dermal cells in the region where a feather primordium will form. It was thought to occur later than the placode stage. However, with better detection techniques (detection of aggregates comprised of a couple of cells versus hundreds of cells), the beginning of its existence has been pushed earlier. Symmetric short feather bud. The short bud is the stage when the base of a feather bud is longer than the long axis of the bud (from the midpoint of the feather base to the tip of the bud). In the early short bud, the bud has radial symmetry. Asymmetric short feather bud. In the later short bud, the bud develops anterior-posterior asymmetry, slanting toward the posterior. The anterior end of a feather bud is defined as the side that forms an obtuse angle with the integument surface, and is the site where the rachis will form. In some feathers, the posterior end sometimes develops into the afterfeather (Fig. 5.2 and 5.5).3 The bud has bilateral symmetry. Short feather buds refer to both short bud stages. Long feather bud denotes the stage when feather buds grow out from the body surface and the long axis of the bud is longer than the base of the bud. Feather filament stage is the time that feather bud epithelia start to form barb ridges. It also occurs when the feather bud starts to invaginate into the dermis to form the feather follicle.

Feather Morphogenesis

63

Short Bud

Long Bud

Ant

Post

Median and Long Axis

Long Axis

Median Axis

Fig. 5.5. Asymmetry formation: or the formation of the A-P axis. When the short feather bud becomes the long feather bud, rather than growing straight (left panel), it becomes slanted. The mechanism can be through differential cell proliferation and movement (middle and right). Because there is more cell proliferation in the posterior bud compartment, we think the mechanism on the right is more likely. The median axis and long axis of the feather bud are defined (see text) to help describe the morphology. Regulators (–) Smaller opening

E=LxPx2

(+) Larger opening

Anagen cycle Cell cycle

Fig. 5.6. Elongation: or the formation of the P-D axis. Following the formation of the feather follicle, epithelial cell proliferation occurs in the collar region (or matrix of the hair) above the dermal papilla. For a unit of area, starting from a fixed number (P) of cells at the beginning an of anagen phase, the more times that cells can divide, the longer the epithelial appendage will be. The time of cell division is decided by the length of anagen period divided by the time required for one cell cycle. The length of the epithelial appendage (E) is the length of a mature epithelial cell (L) times the number of cells stacked proximo-distally. Regulators can modulate the rate which cells require to transit from the proliferation compartment into the postmitotic differentiation compartment. The faster this rate is, the faster the proliferation cell pool will be depleted, and the shorter epithelial will be the appendage.

64

Molecular Basis of Epithelial Appendage Morphogenesis

Feather follicle stage represents the mature feather. It is characterized by a well formed follicular epithelial wall and dermal sheath. The feather in the mature follicle will go through molting changes in the adult bird. In studying feather development, researchers should be aware that there are four growth gradients during feather morphogenesis. The first maturation gradient is across the tract and parallels the order of development. The second maturation gradient is found within a single feather, with distal regions maturing faster than proximal. The third gradient is in terms of the rachis, with barbs closer to the rachis maturing faster than barbs away from the rachis. The fourth gradient is within a barb ridge, with peripheral ridge maturing faster than the central ridge.11 It is important to bear this in mind in interpreting expression data. A properly prepared specimen can consist of different developmental stages. For example, in the spinal tract, the midline buds are more mature than the lateral buds, so that dorsal skin from a stage 37 embryo is composed of four different stages of feather primordia (Fig. 5.3D): primordia in the midline region are at the long bud stage; flanking the midline, primordia are at the short bud stage; further lateral, primordia are at the placode stage; and primordia at the lateral edge of the skin are at the rudiment stage. In the following sections, we will discuss the morphogenetic processes during feather development. In order, we will review: 1. the formation of the tract; 2. the initiation of an individual feather primordium; 3. the formation of the anterior-posterior (A-P) axis; 4. the proximal-distal (P-D) axis; and 5. the formation of barbs and barbules. For each process, we will first describe the phenomenon, review what we know about it, discuss the cellular event(s) that may lead to this morphogenetic process, and then present some known molecular mechanisms involved. It is likely that many key molecules may have not even been identified. Therefore, in each section, we try to define potential molecular players in more general terms, and specific candidates are given only as examples. Future work will put newly identified molecules in appropriate categories and thereby prompt revision of the view presented here. It should be mentioned that another dimension of feather morphogenesis lies in its colors. The many colors and their patterns on the feather make the feather one of the most beautiful organs. The color comes from melanocytes. Chapter 8 discusses molecules involved in the growth control of melanocytes in the avian and mouse.

Macropatterning: Setting Up the Tract Fields To form epithelial appendages, the first step is to establish a field, or a region, on the developing skin where epithelium and mesenchyme become competent to form epithelial appendages. The major tracts include the spinal tract, caudal tract, femoral tract etc. Each tract has its characteristic location, contour, size and number of feathers.3,12 The way different tracts are laid out on the integument surface are termed the “macropattern” (Figs. 5.3 and 5.4).8,13 Although there are events unknown to us, the initiation of a tract is first characterized by the formation of dense dermis. To build the dense dermis, we first need raw material (cells). Where do the cells come from? Using carbon particle labeling and surgical/transplantation experiments, the origin and immigration of the feather mesenchymal cells were traced.14,15 For the trunk feathers, the mesenchymal cells migrate in from the dermatome of the somites. Originally there is a loose fibrous network between the ectoderm and the axial somite/neural tissues. Around embryonic day E3, dermatome cells start to invade, and at around E5 the dense dermis first forms in the midline region above the spinal cord, and

Feather Morphogenesis

Fig. 5.7. Summary of the morphogenetic, cellular and molecular processes in different stages of feather morphogenesis. Stages up to the follicle stages are shown as longitudinal sections. Feather filament stages are shown as cross sections. The morphogenetic processes, cellular processes, and molecular candidates for each process are listed. (-) indicates suppressive action.

65

66

Molecular Basis of Epithelial Appendage Morphogenesis

then expands bilaterally. The dense dermis is about 2.6 nuclei/1000 µm3 and 35-40 µm thick, versus 2 nuclei/1000 µm3 in the dermis of apteric regions. For feathers in the head region, the mesenchymal cells mainly come from the neural crest. For the limb feathers and scales, the mesenchymal cells come from the somatopleura. Throughout the dense dermis, there is also active cell proliferation at this stage. Thus both cell immigration and proliferation contribute to the build up of cell density in the tract region in the initial stage. At about E20 h, the loosely arranged ectoderm begins to form the epithelium by giving rise to the periderm, the basal layer of future epidermis, and the basal lamina. The periderm is present transiently and has protective and other unknown functions. The periderm becomes squamous epithelium from E3 to E6, and eventually sloughs off around E18.16 The epidermis becomes columnar and tightly packed around E6, and gradually becomes stratified.17 Beginning at about E4, the epidermis acquires the ability to respond to dermal signals to form placodes and then skin appendages.7 This competence causes the epidermis within a feather tract to become different from neighboring apteric regions. Within the feather tract, epidermal cells are not fully committed to form the placode or interplacode regions until the long feather bud stage. That is, the placode/interplacode fates are reversible and flexible till the short bud stage.18 To determine where the information for forming a tract is stored, skin epithelia and mesenchyma of different embryonic stages are separated, recombined and cultured. These experiments have shown that the morphological determinants, including the spatial arrangement of the buds and the directionality of sequential formation in the tract, are stored in the mesenchyme. In fact, mesenchyme as early as E2 has received messages specifying it to induce feather tracts.5,7 However, in the early stages, the mesenchymal cells have a certain level of plasticity. For example, implantation of an extra limb bud in the trunk region led to the formation of an ectopic femoral tract that “invaded” into the spinal tract.13 The original spinal tract became asymmetric with a big indentation to accommodate the ectopic femoral tract. This new organization seemed to occur at the expense of the original dermis, which was previously destined to become spinal tract feathers. The remaining spinal tract feather buds’ size and spacing remained the same. Thus, although the bud size and arrangement are stored as intrinsic properties of a specific dermal region, they can be reset before feather bud formation is stabilized. Although epithelia have to be competent to form skin appendages, the origin of the epithelium seems to be a less stringent requirement for skin appendage formation. Epithelia as diverse as apteric epithelium, shank epithelium, foot pad epithelium, cornea, and chorioallantoic membrane, when derived from a competent stage, can respond to feather mesenchyme and form feather buds (reviewed in ref. 7). However, the epithelium does store information directing the orientation of individual feather buds (see below). This should not be confused with the directionality of the tract, which configures the direction of sequential bud formation and is under mesenchymal control. There must be specific molecules expressed in the tract fields that confer competence to the epithelium and mesenchyme to form periodically arranged appendages. We have observed some molecules expressed homogeneously in the dense dermis of the whole tract, but not in the adjacent apteric regions. For example, NCAM is expressed at a basal level in the dense dermis.19 When feather rudiments start to form, NCAM expression becomes enhanced in dermal condensations but decreases in the inter-condensation dermis, thus transforming a homogeneous expression pattern into punctuated patterns. Another example is the signaling molecule, protein kinase C (PKC). Pan-PKC immunoreactivity is homogeneously distributed in the whole dense dermis, and then disappears from each site where a rudiment begins to form.20 Is the presence of NCAM essential to competence? Does the inhibitory action of PKC exert a break in skin appendage formation? The answers to both

Feather Morphogenesis

67

questions are yes, but these are only small parts of the molecular cascades regulating feather development. What constitutes the competence remains to be determined.

Micropatterning: Generation of Feather Primordia Within the Tract With the tract field set, the homogeneous dense dermis and epidermis are transformed into periodically arranged feather primordia in a few days (Fig. 5.4). The arrangement of individual feather buds within the tract is termed the micropattern.8 In vivo, the primordia appear in an exquisite temporal and spatial sequence.21-24 Feather primordia first appear from one end of the primary row and propagate linearly to set up the primary row (or the initial row according to Linsenmayer, 1972).24 Then they propagate unilaterally (e.g., femoral tract) or bilaterally (e.g., spinal tract) to the margin of the tract. By E11, there are approximately 1200 contour feather buds in the chicken.7 Is this sequential order critical for the initiation and spacing of feather primordia? Or, does it just coincide with the maturation gradient? The importance of propagation was emphasized by Sengel (1976)7 who cited experimental evidence that a filter can block the lateral propagation of feather buds, and that glucocorticoid treatment always blocked whole rows (used to indicate the horizontal arrangement from the midline to the lateral edge) of feather buds that were lateral to (or younger than) the directly inhibited ones.7 However, the essentiality of sequentiality for propagation has not been proven. The fact that feather buds can form from aggregates of dissociated epithelial and mesenchymal cells25 suggests that it is not essential. Our recent work also suggests that high cell density alone can initiate feather primordia simultaneously without sequential formation. We suggest that propagation is a more effective, but not essential, mechanism to lay out feather patterns (Jiang et al, in preparation). Is it epidermis or dermis that initiates the feather rudiment? By separating developing epidermis and dermis, it was found that placodes are visible 1-2 rows ahead of the visible dermal condensations.7 This led to the suggestion that the placode may be the initiator. However, the order of appearance depends on the sensitivity of the detection method. A small dermal cell aggregate (say 3-5 cells) may form before placode formation, but it would not be detected as a dermal condensation without sensitive detection methods. Similarly, molecular changes in the epithelium may precede the placode morphology but remain undetectable. While we can continue to push the initiation of the feather rudiment earlier using better methods, it is probably better to answer this question with an “operational definition”, namely, by judging the ability to initiate new primordia. The fact that, upon epithelial-mesenchymal recombination, the old placodes disappear in a few hours, and that new placodes are induced at the sites of previous dermal condensation,18 suggests that the mesenchyme is more likely to be the initiator. Then, what cell processes in the dermis are important? From the dense dermis (about 2.6 nuclei/1000 µm3), feather primordia of about 8,000-15,000 µm2 (approximately 100-140 µm in diameter) emerge in a periodic way. Within the dermal condensations, cell density is about 5.5 nuclei/1000 µm3.26 This differential increase of cell density is probably based on both cell sorting and preferential cell proliferation. Cell proliferation actually stops for about 24 h in the early phase of dermal condensation.26 Thus cell proliferation does not have a role in the early phase of dermal condensation, although it plays important roles in elongation later. It was observed that organized lattices of collagen form before feather primordia become apparent.27 Dermal cells show bipolar fusiform shapes, with the long axis aligned along a plane radiating out from the last formed condensation. These observations led Sengel to suggest that cell adhesion plays a critical role in the patterning of the dermis, and that cells orient themselves according to the organized extracellular matrix.7 The importance of extracellular matrix is further demonstrated in the study that perturbation of

68

Molecular Basis of Epithelial Appendage Morphogenesis

proteoglycan and collagen synthesis leads to disruption of feather development.28,29 Later we observed that dermal cells tend to form aggregates and that antibodies to adhesion molecules NCAM, tenascin, fibronectin, and integrin β1 can perturb dermal condensation formation. These data demonstrate that cell adhesion plays important roles in the formation of dermal condensations.19,30,31 We and others have been tracing upstream regulators for adhesion molecules and have found that several secreted extracellular signaling molecules, such as FGFs, BMPs, Sonic hedgehog, etc., are involved.32-36 Intracellular signaling molecules, such as PKA and PKC are also involved.20 Some of these molecules induce dermal condensation formation, while others suppress it. How these molecules affect dermal condensation formation is discussed further in chapter 13. A fundamental issue in pattern formation is the mechanism regulating the molecular expression that produces periodic feather spacing. Is the periodic feather spacing prepatterned? Is it the manifestation of an endogenous tissue property? Is it timed by an intrinsic clock? Several models for periodic patterning have been proposed37 including chemical reaction-diffusion,38,39 mechanical forces,40 steric hindrance,7,41 etc. Unfortunately the molecular identity of the parameters within the mathematical models have not been identified. Based on our recent experimental data, we present a working model for skin appendage formation in chapter 18. Feathers also can form in cell aggregates without propagation (see Moscona and Moscona, 196525 and our unpublished data). What is the driving force that initiates the formation of feather primordia under those conditions? Our studies on the behavior of mesenchymal cells derived from dermal condensations, as well as from micromass limb bud cells, suggest that mesenchymal cells have an intrinsic ability to form aggregates. The force may be based on cell adhesiveness and random cell movement. Under high cell density conditions, these properties drive cells to randomly form small cellular aggregates. The small aggregates are unstable and reversible. Through competition and support from the epithelia, some small aggregates survive while others get suppressed or merged with other aggregates. During this competitive phase, the boundary of the condensations will gradually sharpen through an autocrine-like positive feedback mechanism (here the unit for autocrine stimulation is the cell aggregate, not a single cell) and lateral inhibition. The final result is the equilibrated state with evenly spaced primordia. DiI labeling showed that when dissociated mesenchymal cells were replated and overlaid with epithelium, the mesenchymal cells randomly redistributed in the primordia or inter-primordia regions of the reappearing feather buds, rather then reverting to their previous locations with predetermination (Jung et al, in preparation). This shows that pattern formation may be achieved based on cell behaviors derived from tissue properties, rather than based on a hardwired program. This is consistent with Dr. Newman’s generic concept of epithelial morphogenesis as discussed in chapter 17 by Newman. In summary, it is proposed that when dermal cells reach a critical density, there is a random process (involving cell adhesion interactions) that produces many initial unstable cell aggregates (the number of these aggregates is more than the final number of buds). Upon reaching a critical size of the cell aggregate, through cell interactions, these aggregates are triggered to send out factors that behave as activators and inhibitors in a reaction-diffusion mechanism. Activators are of short range and make the aggregate grow bigger. Inhibitors are of longer range, diffuse farther and suppress adjacent cells from becoming a part of the feather domain. Through competition and survival, the final pattern that emerges is the equilibrated state of the surviving initiation sites. Successive expression of junction adhesion molecules42 will then stabilize the mature buds. At this stage, we have some candidate

Feather Morphogenesis

69

molecules: FGFs and Shh as the positive factors, BMPs as the negative factors (please see chapter 18). The essence of this view is that the induction of skin appendages is a dynamic process of competition and equilibrium at the level of cellular aggregates, not under a prespecified precise molecular process. More experimental work is required to refine and revise this model.

Anterior-Posterior Axis: Orientation From the feather placode to the early short bud stage, the feather primordia is radially symmetric. Entering the late short bud stage, the radial symmetry shifts to bilateral symmetry with the formation of the A-P axis. The bud gradually slants toward the posterior. For the spinal tract, the A-P orientation is aligned along the cephalic-caudal axis of the body. But this is not always the case. In other tracts, such as the alar tract, the two axes intersect. So it is better to name the anterior and posterior axis based on the individual bud. When and how is this A-P orientation determined? Experiments have been conducted by separating epithelium and mesenchyme of developing skin, and recombining them following 90o or 180o rotation. The results showed that the location is determined by the mesenchymal component, while the orientation is determined by the epithelium.18,43 This is true for the recombinations carried out from stage 29 to about stage 33. Between stage 34 to 35, the orientation activity is gradually shifted to the mesenchyme. In recombination experiments carried out around stage 34, buds around the midline of the original dermis are oriented according to the dermis (because they are more mature), while those at the edges of the dermis are oriented according to the epithelium (because they are still young). Most interestingly, many bi- or triheaded feather buds formed in between.44 These probably occur when the orientation activities from the epithelium and the mesenchyme are about equal. What are the cellular processes that lead to the A-P asymmetry? It could be due to determination of cell fates, or it could be simply explained by differential cell proliferation. In feather buds, we can draw a line from the midpoint of the feather base and perpendicular to the surface of the skin. This line, or the median axis of the bud, separates the bud into anterior and posterior compartments. We can also connect the midpoint of the feather base to the tip of the bud. This line, or the long axis of the bud, separates the bud into anterior and posterior buds. In the symmetric short buds, the median axis and the long axis coincide (Fig. 5.5). In the asymmetric short buds, the two axes become uncoupled. The long axis tilts toward the posterior end and more cells accumulate in the posterior than in the anterior compartment (Fig. 5.5). The cellular mechanism mediating this differential accumulation of cells is not completely understood. Asymmetric distribution of cell proliferation is likely to play a major role because more BrdU accumulates in the posterior bud than in the anterior bud.44,45 Other possible cell processes include uneven cell migration or differential cell death, but we do not have evidence for them yet. Although the molecular cascade that regulates this process is not established, we know that the feather bud exhibits high levels of molecular heterogeneity (see Fig. 13.2). Before morphological asymmetry becomes apparent, we already observe the posterior expression of Wnt7a, Delta-1, Serrate-1, and Notch-1.44 Later, NCAM, tenascin-C, collagen I, acid mucopolysaccharide, and some chondroitin sulfate molecules are localized to the anterior buds, while collagen III and fibronectin are preferentially localized to the posterior buds.31,46,47 We propose that the posterior expression of Wnt7a, Delta, Serrate, and Notch may be involved in establishing preferential proliferation in the posterior compartment. Future work will be required to establish which of these molecules are the cause, and which are the consequence of the A-P asymmetry.

70

Molecular Basis of Epithelial Appendage Morphogenesis

Proximal-Distal Axis: Elongation Having established the boundary of feather primordia and the A-P orientation, the next phase is the elongation of the proximal-distal (P-D) axis. Long feather buds grow by adding cells to the distal as well as the flanking regions. Feathers on the spinal tract reach about 1-1.5 mm at E10. The zone of cell proliferation then gradually shifts toward the base of the buds. In the meantime, the bud base “sinks” (invaginates) into the dermis to become a follicular structure. Gradually, proliferating epithelial cells are organized into a cylindrical conformation immediately above the dermal papilla. From here, cells that are pushed upward contribute to feather elongation. This “cell generator” region is the collar region (Fig. 5.2G), and is approximately 140 µm long.7 Feathers elongate at different rates in different tracts and in different species. For example, they grow faster in the caudal and wing tract. These differences lead to differences in the final lengths of the feathers. The most dramatic example is the tail feathers of the peacock, which can be longer than 1 m and grow at a rate of 8 mm/day.48 In a more typical flight feather, the rate is about 4 mm/day. The range of feather length in a single bird is also very impressive. For example, in an adult parakeet, the shortest mature feather from the head region is 1 mm long, while the longest wing feather of the same bird is 75 mm, a 75-fold difference. What factors regulate the length of the feather? In feathers, new cells are added to the proximal end and differentiate as they progress toward the distal end. The final length of feathers reflects the number of cells along the feather axis multiplied by the average length of cells along the same axis. Cell number is the net sum of cell immigration and proliferation, minus cell emigration and apoptosis. Feathers, similarly to hairs, also have a molting cycle (please see chapter 7). A feather primordium starts with a certain number of epithelial cells in the collar region (probably the transiently amplifying cells discussed by Cotsarelis et al, 1990).4 The feather follicle is a relatively enclosed space, and is not easily accessible for cell immigration or emigration. Cell death does not occur until the catagen phase of the skin appendage. Therefore during the anagen phase, the change of cell number is mainly determined by the number of cell divisions. The number of cell divisions, or the degree of “transient amplifying”, is determined by the duration that cells stay in the proliferative compartment divided by the time required for one cell cycle. The period that a cell can stay in the proliferative phase is influenced by its rate of progression to the differentiation phase and the length of its anagen phase (Fig. 5.6). The molecular factors that regulate the above parameters remain to be determined. However, one example illustrates the linkage between the length of skin appendages and the length of anagen phase. Angora mice have long hairs all over their bodies. Measurements of the hair cycle showed that they have lengthened anagen phase, but the catagen and telogen phase are of the same length.49 It was also shown that FGF-5 knockout transgenic mice have similar long hair phenotypes, and indeed FGF-5 is the mouse Angora locus.50 Therefore FGF-5 is likely to be involved in the transition of anagen to catagen physiologically. Normally, FGF-5 is found in the hair follicle sheath, and several components of FGFs and FGF receptors are found in different regions of the hair follicle.51 By analogy, it is speculated that Angora sheep and rabbits may have molecular defects in FGF-5 or components of the FGF-5 pathway. Similarly, this may contribute to some of the variations in feather length. With recent progress in the molecular control of the cell cycle (a review) and diurnal cycle, the molecular control of the skin appendage cycle is a new and exciting field waiting to be explored (Please see chapter 7). Another regulator is growth control, namely: How many times does a cell in the proliferative compartment (collar) divide before it starts to differentiate? Through the regulation of this transition, the length of the feather can also be regulated. Molecules that accelerate

Feather Morphogenesis

71

this transition will deplete the proliferative compartment quickly and lead to shorter feathers. Molecules that keep cells in the proliferative phase may lead to abnormal new growth. For example, Sonic hedgehog (Shh) is expressed in feathers and is involved in their development.33 Recently, it was shown that K14 promoter-Shh transgenic mice develop basal cell carcinoma that is derived from the hair follicles.52 Thus a balanced molecular control allowing cell proliferation with a steady stream of cell differentiation will lead to feathers with proper length (Fig. 5.6). Many genes involved in new growth, including tumors, are likely to be involved in regulating proliferation and differentiation within the feather. Indeed, feathers are formed from cells with regulated new growth, while tumors are formed by cells with deregulated new growth. During the elongation phase of feather buds, c-Myc, c-Ets, and c-Myb are highly expressed.53 Feedback is an important component of length regulation. One example recently shown in cartilage elongation involves a negative feedback loop of cartilage hypertrophy via Indian hedgehog, parathyroid hormone related protein (PTHRP) and the PTHRPreceptor.54 In summary, elongation is a fundamental process in skin appendage morphogenesis. Other skin appendages, such as tiger claws, elephant tusks and mouse incisors, have shown an enormous elongation potential that makes them powerful weapons. Humans use tools to control their environment, so changes of their physical traits are not as essential for survival. However, in male type alopecia, scalp hairs become small vellus hairs. The possibility of elongation of these tiny hairs to become scalp hairs occupies the minds of many men.

Feather Filament Morphogenesis In the feather follicle, branch morphogenesis occurs above the collar region. The epithelial cells, now 5-6 layers thick, begin to organize into barb ridges, starting from the rachis region (anterior end) and spreading bilaterally (Fig. 5.2). Thus the ridges closer to the rachidial ridge are more mature. The valleys deepen, and epithelial cells lining the valley form the marginal plate. Between the marginal plates are the barb plates with cell proliferation centers located at the tip (toward the center) of the barb ridges. Later cells in the marginal plate die and leave spaces in between barb ridges. This process, plus the continuous presence of cell proliferation in the barb ridges, gives rise to the inter-barb spaces. Later, similar processes take place to form the inter-barbule space and barbules. What is the basis of this epithelial compartmentation that leads to the alternating barb plates and marginal plates? Cells in the collar proliferate and form multi-layered epithelia. The earliest recognizable cellular event is the periodic invagination of this epithelium. This event coincides with the appearance of weak Shh expression in the initial invagination for the presumptive marginal plate.34 As the marginal plate matures, Shh expression becomes more enriched in these epithelia and forms a striking stripe pattern revealed by in situ hybridization (Fig. 5.3D). In the more mature barb ridges, about three ridges away from the initiation of the valley, NCAM starts to appear in the cells situated within the valley. NCAM expression in the marginal plate then extends toward the tip of the barb ridge and gradually intensifies as the marginal plates mature.55 The marginal plate epithelia eventually die and leave space between the barbs (Fig. 5.1). Cells in the barb plate also die, but they leave feather keratins behind, forming the feather proper. The association of Shh and NCAM with the marginal plate probably implies that Shh, a signaling molecule, is involved in setting up the periodic patterning of the barb plate and marginal plate, while NCAM consolidates the boundary of these compartments. Cell death related pathways are then activated in the marginal plate epithelia. The relative lengths of the rachis, barbs, and barbules form a major basis for the variety of feather morphology (Fig. 5.1). Some are radially symmetric (with very short or no rachis),

72

Molecular Basis of Epithelial Appendage Morphogenesis

and others are bilaterally symmetric. Some bilaterally symmetric feathers have a small mirror imaged feather (afterfeather) based on the hyporachis which is 180o opposite to the rachis (Fig. 5.2). Some feathers are fan-shaped, while others are elliptically-shaped. Some feathers are left-right symmetric, and some are left-right asymmetric (with reference to the rachis). Recombination experiments showed that the number of barb ridges are species and region specific. They also showed that the barb pattern is determined by the mesenchyme, but the barbule cell number and morphology are determined by the epithelium.56 The existence of diverse feather morphologies in the same bird provides a model for the study of growth control encompassing epithelial cell proliferation, death and differentiation. A simple heterochrony-like mechanism that regulates these three processes can lead to profound effects on feather morphology.

Perspectives In this chapter we have reviewed how a small piece of presumptive skin can be built into a splendid, branched structure with successive morphogenetic processes (Fig. 5.7). The translation of the one dimensional genomic sequence into the three dimensional structure57 is most dramatic in this example of organ construction. The final shape of a feather depends on the modulation of the morphogenetic processes, and this modulation is the basis of morphological specificity. The distinct morphology of feathers has enabled us to perform a more detailed morphological analysis of epithelial appendages. We can now study the roles of different cellular and molecular processes in the formation of each axis. We hope that the principles described here can be useful in understanding how other epithelial appendages are constructed. It is also hoped that in the future this knowledge may be helpful in developing new ways to manage the repair and regeneration of injured or lost epithelial appendages, and also other organs generated via epithelial-mesenchymal interactions.

Acknowledgment We thank Drs. Jian-fen Lu and Ting-Xin Jiang for help with Figure 5.3. This work is supported by grants from NIH and NSF.

References 1. Gill FB. Ornithology, 2nd edition. Freeman, New York. 1995. 2. Brooks B. On the Wing. Educational Broadcasting Co. 1989:1-33. 3. Lucas AM, Stettenheim PR. Avian Anatomy. Integument. In: Agriculture Handbook 362. Washington DC: Agricultural Research Services, US Department of Agriculture, 1972. 4. Cotsarelis G, Sun TT, Lavker RM. Label-retaining cells reside in the bulge area of pilosebaceous unit: Implications for follicular stem cells, hair cycle, and skin carcinogenesis. Cell 1990; 61:1329-1337. 5. Saunders JW, Gasseling MT. The origin of pattern and feather germ tract specificity. J Exp Zool 1957; 135:503-528. 6. Dhouailly D. Formation of cutaneous appendages in dermo-epidermal interactions between reptiles, birds and mammals. Roux Arch Dev Biol 1975; 177:323-340. 7. Sengel P. Morphogenesis of Skin. Cambridge: Cambridge University Press, 1976. 8. Sengel P. Feather Pattern Development. Ciba Foundation Symposium. 1978; 29:51-70. 9. Chuong CM. The making of a feather: Homeoproteins and retinoids and adhesion molecules. BioEssays 1993; 15:513-521. 10. Widelitz RB, Jiang T-X, Noveen A et al. Molecular histology in skin appendage morphogenesis. J Micro Res & Tech 1997; (in press). 11. Haake AR, Konig G, Sawyer RH. Avian feather development: Relationships between morphogenesis and keratinization. Dev Biol 1984; 106:406-413. 12. Holmes A. The pattern and symmetry of adult plumage units in relation to the order and locus of origin of embryonic papillae. Am J Anat 1935; 56:513-535.

Feather Morphogenesis

73

13. Sengel P. Pattern formation in skin development. Int J Dev Biol 1990; 34:33-50. 14. Mauger A. Role du mesoderme somitique dans le developpement du plumage dorsal chez l’embryon de Poulet. I. Origine, capacites de regulation et determination du mesoderme phumigene. J Embryol Exp Morphol 1972; 28:313-341. 15. Mauger A, Sengel P. La pterule spinale de l’embryon de Poulet: territoire presomptif, arrangement et developpement embryonnaire. Devl Biol 1970; 23:609-633. 16. Parakkal PF, Matoltsy AG. An electron microscope study of developing chick skin. J Ultrastruct Res 1968; 23:403-416. 17. McLoughlin CB. The importance of mesenchymal factors in the differentiation of chick epidermis. I. The differentiation in culture of the isolated epidermis of the embyonic chick and its response to excess vitamin A. J Embryol Exp Morphol 1961a; 9:370-384. 18. Chuong C-M, Widelitz RB, Ting-Berreth S et al. Early events during the regeneration of skin appendages: order of molecular reappearance following epithelial-mesenchymal recombination with rotation. J Invest Dermatol 1996; 107:639-646. 19. Chuong CM, Edelman GM. Expression of cell-adhesion molecules in embryonic induction. I. Morphogenesis of nestling feathers. J Cell Biol 1985a; 101:1009-1026. 20. Noveen A, Jiang TX, Chuong CM. Protein kinase A and protein kinase C modulators have reciprocal effects on mesenchymal condensation during skin appendage morphogenesis. Dev Biol 1995b; 171:677-693. 21. Mayerson PL, Fallon JF. The spatial pattern and temporal sequence in which feather germs arise in the white Leghorn chick embryo. Dev Biol 1985; 109:259-267. 22. Davidson D. The mechanism of feather pattern development in the chick. I. The time of determination of feather position. J Embryol Exp Morphol 1983; 74:245-259. 23. Davidson D. The mechanism of feather pattern development in the chick. II. Control of the sequence of pattern formation. J Expl Morphol 1983b; 30:587-603. 24. Linsenmayer TF. Control of integumentary patterns in the chick. Dev Biol 1972; 27:244-271. 25. Moscona MH, Moscona AA. Control of differentiation in aggregates of embryonic skin cells: suppression of feather morphogenesis by cells from other tissues. Dev Biol 1965; 11:402-423. 26. Wessells NK. Morphology and proliferation during early feather development. Dev Biol 1965; 12:131-153. 27. Stuart ES, Moscona AA. Embryonic morphogenesis: role of fibrous lattice in the development of feathers and feather patterns. Science 1967; 157:947-948. 28. Goetinck PF, Carlone DL. Altered proteoglycan synthesis disrupts feather pattern formation in chick embryonic skin. Dev Biol 1988 127:179-186. 29. Marsh RG, Gallin WJ. Toxic effects of beta-aminopropionitrile treatment on developing chicken skin. J Exp Zool 1994; 268:381-389. 30. Gallin WJ, Chuong CM, Finkel LH, Edelman GM Antibodies to liver cell adhesion molecule perturb inductive interactions and alter feather pattern and structure. Proc Natl Acad Sci USA 1986; 83:8235-8239. 31. Jiang TX, Chuong CM. Mechanism of Feather Morphogenesis: I. Analyses with antibodies to adhesion molecules tenascin, N-CAM and integrin. Dev Biol 1992; 150:82-98. 32. Nohno T, Kawakami Y, Ohuchi H et al. Involvement of the Sonic hedgehog gene in chick feather development. Biochem Biophys Res Commun 1995; 206:33-39. 33. Ting-Berreth SA, Chuong CM. Sonic hedgehog in feather morphogenesis: Induction of mesenchymal condensation and association with cell death. Dev Dyn 1996a; 207:157-170. 34. Jung H-S, Francis-West PH, Widelitz RB et al. Local inhibitory actiom of BMPs and interfeather bud spacing: A model for periodic pattern formation. Dev Biol 1998; 196:11-23. 35. Widelitz RB, Jiang T-X, Noveen A et al. FGF induces new feather buds from developing avian skin. J Invest Dermatol 1996; 107:797-803. 36. Song H, Wang Y, Goetinck PF. Fibroblast growth factor 2 can replace ectodermal signaling for feather development. Proc Natl Acad Sci USA 1996; 93:10246-10249. 37. Held LI Jr. Models for embryonic periodicity. Karger; Basel. 1992. 38. Turing AM. The chemical basis of morphogenesis. Phil Trans R Soc B 1952; 237:37-72.

74

Molecular Basis of Epithelial Appendage Morphogenesis

39. Koch AJ and Meinhardt H. Biological pattern formation: From basic mechanisms to complex structures. Review of Modern Physics 1994; 66:1481-1508. 40. Oster GF, Murray JD, Harris AK. Mechanical aspects of mesenchymal morphogenesis. J Embryol Exp Morphol 1983; 78:83-125. 41. Ede DA. Cell behaviour and embryonic development. Int J Neurosci 1972; 3:165-174. 42. Serras F, Fraser S, Chuong C-M. Asymmetric patterns of gap junctional communication in developing chicken skin. Development 1993; 119:85-96. 43. Novel G. Feather pattern stability and reorganization in cultured skin. J Embryol Exp Morph 1973; 30:605-633. 44. Chen C-WJ, Jung H-S, Jiang T-X et al. Asymmetric expression of Notch/Delta/Serrate is associated with the anterior-posterior axis of feather buds. Dev Biol 1997; 188:181-187. 45. Desbiens X, Turque N, Vandenbunder B. Hydrocortisone perturbs the cell proliferation pattern during feather morphogenesis: Evidence for disturbance of cephalocaudal orientation. Int J Dev Biol 1992; 36:373-380. 46. Mauger A, Demarchez M, Herbage D et al. Immunofluorescent localization of collagen types I and III, and of fibronectin during feather morphogenesis in the chick embryo. Dev Biol 1982; 94:93-105. 47. Kitamura K. Distribution of endogenous beta-galactoside-specific lectin, fibronectin and type I and III collagens during dermal condensation in chick embryos. J Embryol Exp Morphol 1981; 65:41-56. 48. Brun R. Beitrag zur Kenntnis der Dynamik im Federkeim. Rev Suisse Zool 1968; 75:1056-1063. 49. Sundberg JP, Rourk MH, Boggess D, Hogan ME, Sundberg BA. Bertolino AP. Angora mouse mutation: Altered hair cycle, follicular dystrophy, phenotypic maintenance of skin grafts, and changes in keratin. Vertinary Pathology (in press). 50. Hebert JM, Rosenquist T, Gotz J et al. FGF5 as a regulator of the hair growth cycle: Evidence from targeted and spontaneous mutations. Cell 1994; 78:1017-1025. 51. Rosenquist TA, Martin GR. Fibroblast growth factor signalling in the hair growth cycle: expression of the fibroblast growth factor receptor and ligand genes in the murine hair follicle. Dev Dyn 1996; 205:379-386. 52. Oro AE, Higgins KM, Hu Z et al. Basal cell carcinomas in mice overexpressing Sonic hedgehog. Science 1997; 276:817-821. 53. Desbiens X, Queva C, Jaffredo T et al. The rlationship between cell proliferation and the transcription of the nuclear oncogenes c-myc, c-myb and c-ets-1 during feather morphogenesis in the chick embryo. Development 1991; 111:699-713. 54. Vortkamp A, Lee K, Lanske B et al. Regulation of rate of cartilage differentiation by Indian hedgehog and PTH-related protein. Science 1996; 273:613-622. 55. Chuong CM, Edelman GM. Expression of cell adhesion molecules in embryonic induction. II. Morphogenesis of adult feathers. J Cell Biol 1985b; 101:1027-1043. 56. Sengel P, Dhouailly D. Differenciation en greffe chorioallantoidienne de chimeres interspecifiques de peau embryonnaire de poulet et de canard. CR Acad Sci, serie D 1966; 263:601-604. 57. Edelman GM, Morphoregulation. Dev Dynamics 1992; 193:2-10.

CHAPTER 6

Principles of Hair Follicle Morphogenesis Michael Philpott and Ralf Paus

Introduction

T

he hair follicle is the most prominent cutaneous mini-organ and one of the defining features of mammalian species. Only mammals display hair follicles, although not every mammalian species does so; whales, for example, are devoid of hair. As discussed in chapters 5 and 19, hair follicle morphogenesis follows ancient, evolutionarily highly conserved, strategies for organ development that involve a closely coordinated series of bidirectional epithelial-mesenchymal interactions which dictate all stages of induction and morphogenesis. These interactions display striking similarities between seemingly unrelated systems, such as the development of fish and reptile scales, feathers, nails, claws, hooves, antlers, teeth, limb buds, mammary glands—and hairs.1-5 While feathers are thought to have evolved from reptile scales,6 the evolutionary ancestry of hair follicles is not quite clear. There is general agreement that hair is not the homologue of feather, tooth, nail and scale. In fact, the first hair precursors may have arisen as part of sensory structures between the scales of reptiles. Resemblences of this can still be appreciated in the tail region of rodents, where groups of hair follicles arise in the hinge region of the reptile-like scales of rodent tail skin.1,3,6 Thus, the “sensory” tylotrich hair follicle of rodent skin, with its associated, innervated epidermal cluster of Merkel cells (Pinkus’ Haarscheibe or tactile disc),7 may have been a critical step in modern hair follicle evolution. This evolutionary descendence from sensory structures may also explain why all hair follicles are exquisitely innervated, and why some hair follicles, most notably whisker follicles (vibrissae) have developed into high-sensitivity sensory organs.8,9 Ever since the early days of modern scientific interest in skin, the development of hair follicles has fascinated physicians, anatomists and zoologists alike for the precise choreography of morphological events in both the skin epithelium and the skin mesenchyme that culminates in the development of this fairly complex mini-organ. Even today, one can only marvel at the systematic and careful analyses of follicle anatomy and morphogenesis that have been left to us by some of the founding fathers of skin biology, perhaps most notably by Koelliker,10 Unna,11 Stohr,12 and Dry13 as well as by Felix and Hermann Pinkus.14-18 In fact, almost forty years after Hermann Pinkus’ detailed histological analysis on human embryonal hair follicle development,16 it is hard to improve on these phenomenological studies, whose insightful and often visionary interpretations have anticipated many key concepts which have become incorporated into the mainstream of modern hair biology. Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

76

Molecular Basis of Epithelial Appendage Morphogenesis

Hair follicle morphogenesis can be approached from several different angles: one can focus on where and when follicles develop, why they develop in a specific location and not another, how they develop and also why. Also of significant interest is the variation in follicle type (e.g., pelage hair, vibrissae, cilia, genital hairs) and the sequence in which they occur during the ontogeny of mammalian development. Moreover, one can also ask how the development of an individual hair follicle is coordinated with that of others. For example, in sheep, a trio of one central and two lateral follicles develops in the direct vicinity of a large primary hair follicle, and remnants of such developmentally (and even functionally?) linked hair follicle collectives may be found even in human skin.3,18 While all of these different angles pose biologically most intriguing—and essentially unsolved—problems, for the purpose of this chapter we shall adopt a narrow focus, centered on the morphological features and the possible molecular controls of pelage hair follicle development in man and the mouse.

Functional Anatomy of the Hair Follicle At the beginning of this account of hair follicle morphogenesis, it is useful to familiarize oneself briefly with its final product, the mature anagen hair follicle, so as to better appreciate the developmental steps that have to be taken in order to assemble this multicylindric epithelial-mesenchymal interaction unit. In essence, this “perfect microcosmic structure” (Rothman) can be viewed as a high output fiber-production factory, whose complete production unit, the epithelial hair bulb, is periodically constructed and deconstructed,19 as if it were responding to some as yet unknown changes in the biological “demand” for pigmented hair shafts. Several concentrically arranged epithelial cell layers, each characterized by its own differentiation program and special biochemical, mechanical and secretory properties, collaborate with hair bulb melanocytes to generate a pigmented hair shaft. All this is apparently under the command of a highly specialized fibroblast population that retains its inductive properties even in the adult mammalian organism, the cells of the dermal papilla (DP).19-22 Moreover, in view of the importance of neuroectoderm-derived follicular melanocytes for hair shaft pigmentation, of the elaborate networks of sensory and autonomic follicle innervation, and the increasing evidence in support of neural mechanisms of hair growth control, the hair follicle may indeed best be viewed as an epithelial-mesenchymal-neuroectodermal interaction unit.8,9,23 The major structural components of the anagen hair follicle include the dermally derived papilla (DP) and connective tissue sheath (CTS), the epithelial hair matrix (HM), including the germinative cells (GE), and the outer root sheath (ORS). Hair growth is effected by proliferation of epithelial germinative cells located in the follicle bulb, which are believed to be under the control of the dermal papilla.21,22,24 Germinative cell division gives rise to matrix cells which undergo lineage restricted differentiation to give rise to at least six concentric rings of cells which form the cortex and cuticle of the hair fiber as well as the cuticle, Huxley’s and Henle’s layers of the inner root sheath. Since the final stage of hair follicle morphogenesis morphologically resembles a mature anagen hair follicle, Figures 6.1c and 6.2 [Stage 8 ] illustrate the basic architecture of the anagen hair follicle. Hair growth (anagen) is followed by apoptosis-driven regression (catagen) during which the epithelial cells undergo apoptosis and tissue remodeling to form a small resting (telogen) follicle that re-enters anagen. Thus the hair growth cycle appears to mimic certain aspects of the fetal morphogenesis and cytodifferentiation of the hair follicle and the anagen stage of the hair growth cycle would appear to recapitulate, at least in part, embryology (see chapter 7). In humans the hair growth cycle is asynchronous (“mosaic”), and each individual

Principles of Hair Follicle Morphogenesis

77

hair follicle undergoes its own internal cycle and appears to be largely independent of adjacent follicles. In other mammals, however, including rodents, the hair growth cycle is synchronized and follows a specific pattern of growth and regression, particularly in the early postnatal period. The signaling mechanisms that are involved in both the embryonic development of the hair follicle and in its subsequent growth and cycling in the adult are still poorly understood. However, development of the mammalian tooth and the avian feather have been widely studied, and their mechanisms of embryonal induction appear to be at least in part analogous to that of the hair follicle. Subsequent development of hair follicle, tooth and feather are, then, driven and controlled by a series of precisely choreographed, reciprocal epithelial-mesenchymal interactions involving, e.g., secreted growth factors, differentially expressed growth factor receptors and transcription factors, adhesion molecules and changes in the extracellular matrix milieu, with similar molecular themes of morphogenesis control appearing during the development of all three different appendages25,26 (also see introductory chapter 1). Mice have long provided exceptionally instructive hair research models, and most basic principles of hair follicle biology and pharmacology were first established in the murine system.4,19,20,27-31 Furthermore, the molecular basis of many spontaneously arisen mouse mutations with hair growth abnormalities, that have been studied by hair researchers for decades, is gradually being elucidated.31-34 Typically, it is only after recognition and molecular definition of a mouse mutation with a specific hair phenotype that a human analog is discovered, as is the recent landmark discovery of a human analog of the mouse hairless gene. Together with the ever-increasing number of novel mouse mutants where the genetically engineered knockout or overexpression of a defined gene-product is associated with a hair phenotype (e.g., Angora, waved-1 and waved-2), this has allowed unparalleled insights into the molecular controls of hair growth. Unfortunately, however, this has not yet stimulated the synthesis of truly convincing scenarios for the molecular basis of hair follicle induction and morphogenesis that would reach the level achieved in the fields of feather and tooth morphogenesis (see chapters 5, 9 and 14), and many current molecular concepts of hair follicle morphogenesis rely heavily on drawing parallels to tooth and feather development.

Why Hair Follicle Development Matters As delineated before (see chapter 15), hair follicle morphogenesis offers a perfect, easily dissectable model system for dissecting the molecular controls of induction, pattern formation and morphogenesis in general. Hair follicle development, therefore, is not only of the utmost interest to the wool industry and to physicians dealing with congenital hair growth abnormalities, but also to a wide range of biologists with an interest in developmental systems and epithelial-mesenchymal interactions. Embryologically, the hair follicle is the result of complex, bidirectional epidermal-mesenchymal interactions,4 into which regulatory signals from cutaneous nerves as well as from the endocrine and immune system may feed.23 Knowledge of the molecular controls of follicle morphogenesis may enable us to induce the de novo formation of hair follicles in adult human skin, where these miniorgans have been lost as a consequence of disease. That this is not an entirely utopian goal is documented by the fact that experimental hair follicle neogenesis in mature mammalian skin has already been reported in several rodent species long after birth22,35-37 and that some dermatofibromas in aging human skin are associated with de novo development of immature, folliculoid structures.18

78

Molecular Basis of Epithelial Appendage Morphogenesis

The follicle is unique in that some of these developmental interactions appear to persist in the adult follicle, resulting in a cyclical pattern of growth (anagen), regression (catagen) and relative “rest”19,20 (see chapter 7). It has long been felt, therefore, that a deeper understanding of the molecular controls of hair follicle development may eventually help to manage common hair growth disorders, many of which can be understood as disorders of hair follicle cycling.19 Based on striking morphological similarities between embryonal follicle development and the cyclic telogen-anagen transformation of the hair follicle, it is a widely held concept that the molecular communication that drives follicle induction and subsequent morphogenesis is recapitulated, at least in part, when a resting (telogen) follicle enters into the growth stage of the hair cycle (anagen). If this is true, data on hair follicle development may help to design novel strategies for more efficient induction or suppression of anagen wherever this is clinically desired, such as in cases of telogen effluvium, androgenetic alopecia, hirsutism and hypertrichosis.19 However, it remains to be rigorously tested whether such simplistic analogies between the controls of follicle morphogenesis and anagen development in adult human skin are permissible. Certainly, one needs to consider that the molecular mechanisms of induction, pattern formation, and organ morphogenesis may be restricted to a narrow developmental window. However, the fact that hair follicle neogenesis apparently does occur under certain conditions even in later postnatal life34,36,37 raises optimism that some of these developmental controls are still operative in adult skin and that they may be exploitable for inducing or suppressing anagen development in the adult.

Morphology of Hair Follicle Development In order to call attention to the—often unduly neglected—foundations of the field, it is appropriate to begin our account of hair follicle morphogenesis with a reproduction of Stohr’s magnificent 1903 lithographs, (Fig. 6.1) and to summarize the morphological principles of hair follicle development on the basis of H. Pinkus’ magisterial 1958 account of human fetal hair follicle development. This is complemented by phenomenological data obtained by later researchers. We then switch to a delineation and discussion of murine hair follicle morphogenesis, providing the reader with a schematized system that allows easy recognition and classification of the various stages of murine hair follicle morphogenesis. The murine system has generated the most incisive insights into the molecular controls of hair follicle development. Drawing on recent molecular data from the murine system, and on analogies to the— much better defined—molecular biology of tooth and feather development, we shall finally sketch potential epithelial-mesenchymal signaling pathways that may underlie the control of hair follicle induction and morphogenesis, and define open key questions.

Human Hair Follicle Morphogenesis The different stages of human hair follicle development are summarized in Table 6.1 and illustrated in Figure 6.1. According to F. and H. Pinkus,14-16 the following stages are traversed in chronological order by the developing follicle epithelium: pre-germ (“Haarvorkeim”); hair germ (“Haarkeim”); hair peg (“Haarzapfen”); bulbous peg (“Bulbuszapfen”); hair bulb differentiation with the gradual appearance of hair matrix (HM), inner and outer root sheaths (IRS, ORS), pigmented hair shaft, bulge (“Wulst”), isthmus, and infundibulum. Sebaceous gland, apocrine gland, and arrector pili muscle develop concommittantly. In humans the early embryonic skin (<60 days EGA-[estimated gestational age]) is composed of a simple epithelium derived from the surface ectoderm, consisting of a basal layer and an outer periderm; and a dermis derived from both the somatopleuric layer of the lateral plate mesoderm and from the dermatomal divisions of the somites. At around day 60

Principles of Hair Follicle Morphogenesis

79

Table 6.1. Stages of human and rodent hair follicle development

Stage Prefollicle Follicle Initiation Pregerm Hair Germ Hair Peg Bulbous Hair Peg Lanugo Hair Follicle

Human§ EGA

Pelage

8-9 weeks

12-13 days

10-11 days

10-11 weeks 11-12 weeks 12-15 weeks 15-18 weeks 18 weeks to birth

14 days 15 days 16 days 19 days ***

12 days 13 days 14 days 14-15 days ***

§ After Kaplan and Holbrook54 # After from Davidson and Hardy55

Rodent (dpc)# Whisker

(EGA: estimated gestational age) dpc: days post coitum)

EGA the embryonic skin stratifies to produce an intermediate layer which serves as the precursor of the outer differentiating layers of the mature epidermis.38 The human hair follicle is derived from both the epidermis and dermis and is composed of epithelial components, the matrix and outer root sheath, and dermal components, the dermal papilla and connective tissue sheath. Hair follicle development in the human embryo is first seen at around day 80 EGA with the formation of hair germs in the region of the eyebrow and then the scalp; further hair germs then develop over the rest of the body in a cephalo-caudal direction. Thus by 15 weeks EGA hair follicles have reached the hair peg stage on the upper body but have not begun to develop on the lower body, where the hair germ stage only appears around 17 weeks EGA.39 Positioning of hair germs within the skin appears to be very tightly regulated, and follows a specific distribution pattern which seems to be dependent upon the body region. However, once the site of hair follicle development has been established, each individual follicle within that specific region develops independently of its surrounding follicles, in an asynchronous fashion.18 The first stages of hair follicle development in human skin are characterized by the formation of the prehair germ stage, defined by Pinkus16 as a crowding of nuclei in the basal layer of the epidermis. This crowding of nuclei was later shown by the light and electron microscopical studies of Breathnach and Smith,40 Hashimoto41 and Carlsen42 to be the result of basal epidermal cells forming tightly packed placodes. These placodes then become associated with aggregates of nonproliferative mesenchymal cells43 which form within the dermis and, along with the epithelial placode, give rise to the definitive hair germ (Fig. 6.1a).40,41 Aggregation of mesenchymal cells below the pregerm stage appears to be essential for subsequent hair follicle development and, if these mesenchymal aggregates fail to develop or if their formation is disrupted, then further follicle development does not proceed. Thus histologically it would appear histologically that the initial instruction to form a follicle resides in the epidermis with the formation of the epidermal placode, and that the control of subsequent follicle development is then transferred to the mesenchyme. Once the hair germ has formed, the embryonic follicles grow down into the dermis to form the hair peg stage. The advancing edge of the hair peg forms a broad club of columnar epithelial cells organized along the long axis of the follicle. In contrast, the peripheral cells of the hair peg form radial to the long axis. The center of the hair peg is composed of small, more rounded cells,44 and the point where the hair peg interfaces with the epidermis is characterized by a column of cells which extend back into the epidermis to form the anlage of the hair canal through which the keratinized hair fiber will erupt to the surface of the

80

Molecular Basis of Epithelial Appendage Morphogenesis

skin.39 Though hair follicle formation is traditionally summarized as an epidermal invagination, Pinkus has correctly pointed out that the downwards-moving epithelial cell column is, in fact, largely generated by mitosis, not by an invagination-like migration of epidermal cells into the dermis.16 As the hair peg develops, it becomes enveloped in a sheath of mesenchymal cells 2-3 layers thick. These mesenchymal cells appear to be derived from the mesenchymal aggregate and in the fully developed hair follicle will form the connective tissue sheath. Further downgrowth of the hair peg results in the epithelial cells at the base of the follicle broadening out to encompass the underlying mesenchymal aggregate, thus forming the presumptive dermal papilla of the hair follicle and giving rise to the bulbous peg stage of development. During the development of the bulbous hair peg, two or sometimes three swellings appear at the side of the follicle. The uppermost swelling develops into the sebaceous gland and the lower most forms the bulge, which acts as the point of attachment of the arrector pili muscle and also contains hair follicle stem cells.30 In embryonic human hair follicles, the epithelial cells of the bulge are smaller and more closely packed than surrounding cells. Moreover, the bulge of embryonic human hair follicles contains keratin K19 positive cells.45 This is important, as keratin K19 has been suggested as a possible marker for stem cells in other tissues.46 In adult hair follicles, clearly defined bulges are usually best seen in follicles from the eyelash and can only be found with difficulty in follicles from other body sites. The third swelling, which if present forms above the developing sebaceous gland, gives rise to the apocrine sweat gland, which plays an important role in body odor. However, development of the apocrine gland is usually restricted to the follicles of the armpit and perianal regions of the body is generally absent from other body sites. Once the follicle bulb and sebaceous gland have developed (Fig. 6.1b,c), the germinative epithelial cells in the

Fig. 6.1. Human hair follicle development. From: The Biology of Hair Growth. With kind permission of Academic Press.

Principles of Hair Follicle Morphogenesis

Fig. 6.1b.

Fig. 6.1c.

81

82

Molecular Basis of Epithelial Appendage Morphogenesis

follicle bulb proliferate to give rise to a population of transitory amplifying cells which migrate upwards within the hair follicle and undergo lineage restricted terminal differentiation giving rise to several concentric cones of cells. The inner most layers of cells form the component structures of the hair fiber, the cortex and cuticle, whilst the outer layers form structures of the inner root sheath, namely the cuticle of the IRS as well as Huxley’s and Henle’s layers.47 Keratinization within the hair cone is reported to proceed as early as the 15th week EGA and occurs first in the cells that form the outer most layer and tip of the hair cone.48 These cells are destined to form Henle’s layer of the IRS and their terminal differentiation is characterized by, first, the accumulation of trichohyalin granules and then keratin filaments and dense cornified material. The transition to fully hardened, keratinized tissue proceeds very abruptly and is not fully understood. Huxley’s layer is the next to keratinize, because this proceeds slightly later than Henle’s layer and because germinative cells in the follicle bulb are actively dividing and generating new daughter cells which push the hair cone upwards. Then keratinization of Huxley’s layer proceeds more distally within the follicle than that of Henle’s layer. As a result of increased cell proliferation in the germinative cells of the hair follicle bulb, the hair fiber moves upwards within the follicle and keratinizes. As the keratinized hair fiber moves towards the surface of the skin, it enters the central hair canal of the follicle. The hair canal consists of a sub-epidermal portion composed of the isthmus and infundibulum and an epidermal portion, and is formed from a central cord of ‘core’ cells which stem from the developing hair follicle and project upwards into the epidermis.44,49 The exact method by which the hair canal is formed is still uncertain; however, it would appear that in the isthmus and infundibulum of the hair follicle the ‘core’ cells of the presumptive hair canal do not fully keratinize.39,44,49 Instead they appear to undergo a programmed cell death that is characteristic of apoptosis.50 The hair canal is then bounded by 2-5 layers of keratinized IRS cells which themselves desquamate at the level of the sebaceous gland, freeing the hair fiber into the intraepidermal hair canal. The ‘core’ cells of the intraepidermal hair canal would appear to keratinize and desquamate: this blocks the hair canal, but is removed by the hair fiber as it erupts at the surface of the skin. Emergent hair fibers can usually be seen by 17 weeks EGA on the brow, and are visible over the entire scalp by 18-21 weeks.39,51 Emergence of hair then spreads in a cephalo-caudal direction52 until the entire body of the fetus is covered by lanugo hair. This hair is fine and poorly pigmented and is usually shed at 7-8 months in utero as the hair follicles proceed through their first hair cycle.53 On the scalp, shedding of hair occurs in a wave from the frontal to parietal region. The follicles of the occipital region remain in anagen until birth when they also cycle and the hair fiber is shed. This difference in cycling between the frontal and parietal regions of the scalp and the occipital region are interesting, because of the possible difference in embryonic origin of the dermis in these two regions. In the frontal/parietal regions of the scalp the dermis is derived from the neural crest, whereas the occipital region is derived from the lateral plate mesoderm. Following the shedding of the first lanugo coat a second normally grows, which like the first consists of fine, poorly pigmented fibers. This second coat is usually shed shortly after birth and is followed by the growth of fine medullated vellus hair fibers.53 Terminal hairs grow on the scalp, eyelashes and eyebrows. During fetal life, all of the scalp hair follicles are synchronized in the same stage of the growth cycle. This synchronized pattern of hair growth is, however, lost during the first year and follicles assume an asynchronous, mosaic pattern of growth.18

Principles of Hair Follicle Morphogenesis

83

Murine Hair Follicle Morphogenesis Murine hair follicle morphogenesis basically recapitulates the main architectural themes of human hair follicle development delineated above. It, therefore, offers an ideal model system for dissecting the molecular controls of hair follicle development, since it is much more amenable to experimental manipulation than the human follicle. Also, many pelage hair follicles in mice develop very late, i.e., in the perinatal period.13 Even in the skin of newborn mice, all stages of hair follicle morphogenesis can still be encountered and conveniently subjected to analysis and experimental manipulations.23 It is, therefore, worthwhile to familiarize oneself with the basic criteria that allow one to recognize and classify defined stages of hair follicle development in mice. To this end, one can employ a simplified, schematized system for staging follicle morphogenesis, using the development of pelage follicles in the back skin of neonatal C57BL/6 mice as an example.23 This classification system distinguishes eight major stages of hair follicle morphogenesis, in slight modification of the developmental stages suggested by Hardy (Fig. 6.2).4,55 Most of these developmental stages can be identified in simple H&E-stained routine sections, on the basis of morphology alone. Further help in follicle staging can be recruited if one assesses alkaline phosphatase activity (AP) by enzyme histochemistry so as to visualize the exact location and shape of the strongly AP+ dermal papilla.56 Additional information for classifying follicle development can be generated by the immunohistological demarcation of interleukin-1 type I receptor (IL-1RI) and of transforming growth factor β type II receptor (TGF-βRII) immunoreactivity as useful markers for follicle keratinocytes.23,57 When considering the development of murine hair follicles, it is also important to remember that the coat or pelage consists of a number of different hair types.32 The major pelage hair types include the guard hair which makes up approximately 2% of the coat and consists of long straight hair fibers that protrude above the surface of the coat. Awls are straight hair fibers that are usually only half the length of the guard hair and make up 28% of the pelage. Auchenes are similar to awls, but have a single bend in the hair fiber while zigzags have more than one bend in the fiber and make up 70% of the fibers in the pelage. In addition to the pelage fibers we must also consider the vibrissa or whisker, which is produced by a large innervated follicle surrounded by a large blood sinus and serves a sensory function.

Principles of Hair Follicle Induction As we have seen, the earliest histological sign of hair follicle development is the thickening of the epidermis to form a placode. Following formation of the placode, mesenchymal cells aggregate beneath the placode, and skin appendage induction, morphogenesis and finally cytodifferentiation occur. Formation of the epithelial placode and subsequent mesenchymal aggregation also occurs during tooth and feather development and suggests that the initial signal for the development of the hair follicle, tooth and feather resides in the epithelium, and that once the developing appendage has reached the epithelial bud stage, at E12-13, then subsequent control of development moves to the condensing mesenchyme.58 However, experiments using heterotopic recombination of undifferentiated mouse skin suggest that hair follicle morphogenesis may first reside in the specialized dermal fibroblasts condesation that later on develops into the dermal papilla of the hair follicle. The following sequence of events may result in the specification for a hair follicle to develop4 (also see Fig. 15.5): 1. First dermal message—‘Make an appendage.’ 2. Epidermal message—‘Mesenchymal cells aggregate.’ 3. Second dermal message—‘Make a hair follicle.’

Fig. 6.2. Summary of murine hair development. Different rows represent different stages of hair development. For each row, the first column is schematic drawings of the particular stage of follicle development, with all necessary anatomical details. The second column shows a list of the criteria for staging the developing hair follicle. The third column contains two or three pictures showing the features, as described in the second column. The labels in the pictures refer to the criteria. Immunohistological stains used: alkaline phosphatase (AP); transforming growth factor-β receptor type II (TGF-βIIR); interleukin 1 receptor type I (IL-1RI). The figure shows stages 0-8 of hair follicle morphogenesis (stage classification modified after Hardy4) in nionatal C57B6/ 6 mice. Modified after Paus et al, submitted. (Figure continues on following three pages.)

84 Molecular Basis of Epithelial Appendage Morphogenesis

Principles of Hair Follicle Morphogenesis

85

86

Molecular Basis of Epithelial Appendage Morphogenesis

Principles of Hair Follicle Morphogenesis

87

88

Molecular Basis of Epithelial Appendage Morphogenesis

From the recombination experiments, it would appear that the first dermal message is not species-specific and that mouse dermis can initiate placode formation in chick skin and scale placodes in lizard skin.4 However, the epidermal message to make a dermal papilla appears to be species-specific, since only mouse mesenchymal cells will respond to signals from mouse epidermis.4 Once the mesenchymal cells have formed an aggregate under the epithelial placode, subsequent tissue morphogenesis and cytodifferentiation to form a hair follicle appear to reside in the dermis. The importance of the epidermis in formation of the mesenchymal aggregate has been further demonstrated by the experiments of Hirai et al.59,60 These authors have shown that when embryonic skin from E13 mice is maintained with blocking antibodies to both E- and P-cadherin, epithelial morphogenesis was disrupted, and mesenchymal aggregates failed to develop. Analysis of E- and P-cadherin in developing follicles showed that these adhesion molecules were expressed in the epithelium and not in the mesenchyme. This suggests that adhesion molecules expressed by the epithelial cells are topobiologically important in specifying formation of the dermal papilla (see also chapter 15).

Signaling Events in Hair Follicle Development Today, we are faced with an ever-increasing list of candidate molecules implicated in the control of hair follicle morphology. Table 6.2 provides a tentative list of candidates. The relative functional importance and the hierarchy of these signals has yet to be clarified. For the potential role of adhesion molecules in the control of follicle morphogenesis, which will not be dealt with in the current chapter, see chapter 15. Many of these candidate signals are assumed to be involved in the control of hair follicle development, judging from their peculiar expression patterns at the gene or protein level during various stages of follicle morphogenesis. This will be exemplified by a short account of the expression of selected signaling molecules during human hair follicle development (see next section). However, as this account will show, such phenomenological work tells us fairly little about the relative functional importance of the molecule in question, even if it is differentially expressed during the earliest developmental stages. One never knows whether there isn’t another, as yet unrecognized up-stream signal that ultimately controls this differential expression pattern and is, therefore, the primary force driving follicle induction and morphogenesis. Fortunately, there is now a very large number of spontaneous32,33 or experimentally induced (see above) mouse mutations available for study, which show abnormalities in hair follicle development. These offer the chance to probe the relative functional importance of a defined gene and its product for selected stages of hair follicle induction and morphogenesis. In fact, it was only with the help of such mouse mutants that specific candidates for the role of key upstream signals in the control of hair development can now be listed. Given how much the hair follicle depends on highly controlled and delicately balanced epithelial-mesenchymal-neuroectodermal interactions for its orderly development, the large number of genes whose functional knockout or overexpression causes severe abnormalities (Table 6.3) in this complex microcosmos is not surprising. The availability of these mutants (only some of which can be discussed here due to space limitations) has aided greatly in the endeavor to decipher the molecular controls of hair follicle morphogenesis. A section describing a short account of recent insights into the molecular controls of murine hair follicle development obtained from the analysis of such mutants is presented.

Principles of Hair Follicle Morphogenesis

89

Table 6.2. Genes expressed in early hair follicle development Molecule Homeobox genes HoxC8 HoxD9 HoxD11 HoxD13 Msx-1 Msx-2 Other transcription factors Lef1 Whn Id 1-3 Id 4 M-Twist

Localization

Epithelium70 Epithelial cells of hair bud70 Epithelial cells of hair bud70 Late expression in matrix cells70 Hair follicle mesenchyme71,72,73 Epithelial placode. Matrix of adult hair follicle72,73 Pluripotent ectoderm, epithelium of developing follicle, mesenchymal aggregate and DP of adult hair follicle118,119 Epithelium of hair follicle bud35,120 Mesenchymal aggregate, CTS, ORS and DP124 IRS124 Dermal papilla. Decreases during development123

Signaling and adhesion molecules Amphiregulin Bulbous hair peg130 BRCA I Epithelial placode and hair peg131 BMP-2A Ectoderm, epithelial placode, developing cortex and IRS. In adult follicle precortical cells33 BMP-4 Transient expression in hair germ mesenchyme86,87 BMP-3 Weak staining DP, ORS and IRS89 BMP-7 ORS and IRS89 E-Cadherin Upper follicle positive, developing matrix negative59 P-Cadherin Epithelial placode59 Clusterin/TRPM-2 Epithelium of developing follicle132 α1 Connexin Hair germ. Later expression in IRS, ORS and matrix133 α2 Connexin Hair cone and ORS133 CD44 Mesenchymal condensation134 I-CAM Transient expression in outer layer of epithelial cells of hair germ61 Epimorphin Mesenchymal condensation60 EGFR Epithelial placode negative. Basal layer of hair peg positive63,92 FGF-1 (aFGF) Epithelial placode112 FGF-2 (bFGF) DP of bulbous peg112 FGF-7 (KGF) DP113 FGFR1 DP of developing follicle109 FGFR2 Basal layer of epidermis developing follicle epithelium Later expression in both follicular epithelium and DP109 KGFR Epithelium of developing hair follicle113 Hedgehog Epithelial placode before condensation of the mesenchyme90,91,135,136 (continued on next page)

Molecular Basis of Epithelial Appendage Morphogenesis

90

Table 6.2 (continued). Molecule

Localization

Signaling and adhesion molecules (continued) Patched IGF-I c-Met Midkine NGF NGFR Pleiotrophin PDGF-A PDGF-B PDGFRα PDGFRβ TGF-β1 TGF-β2 TGF-β3 TGF-βIR TGF-βIIR Serrate Notch p53 Neurotrophins Retinoic acid receptors RAR α RAR γ

Epithelial placode adjacent to hedgehog90 Mesenchyme of developing hair follicle137 Outer cells of epithelial placode138 Epithelium and mesenchyme of developing follicle125 Epithelium and developing hair follicle139 DP, epithelial bulge, basal cells of ORS61,140 Epithelium of developing follicle. Mesenchyme negative125 Outer epithelial cells of hair germ61 Inner core cells of hair germ61 Dermal condensation61 Dermal condensation61 Epithelium of hair follicle82 Mesenchyme and epithelium81,82 Epithelium82 Epithelium including placode, faint staining of DP23 Focal expression in epithelium prior to placode formation. Mesenchyme negative23 Epithelium. Developing follicle and mesenchyme126 Hair bud epithelium127,128,129 Hair germ epithelium141 Follicle epithelium and mesenchyme142 Mesenchymal aggregate76 Epithelial placode and mesenchymal aggregate76

Extracellular matrix and proteoglycans Collagen I, III and V Absent or downregulated in mesenchymal aggregate54,61 Collagen IV and VII BMZ of epidermis and hair germ54,61 Hyaluronic acid receptor Mesenchymal condensation Syndecan-1 Epithelium and mesenchyme of developing follicle125 Tenascin Upregulation in BMZ of placode and developing follicle54 Perlecan DP positive61 C-S-PG Upregulated in BMZ of developing follicle61 Abbreviations: BMP, bone morphogenic protein; BMZ, basement membrane zone; C-S-PG, chondroitin sulfate proteoglycan; CTS, connective tissue sheath; DP, dermal papilla; EGFR, epidermal growth factor receptor; FGF fibroblast growth factor; FGFR, fibroblast growth factor receptor; KGF, keratinocyte growth factor; KGFR, keratinocyte growth factor receptor; IGF-I, insulin like growth factor-I; IRS, inner root sheath; NGF, nerve growth factor; NGFR, nerve growth factor receptor; ORS, outer root sheath; PDGF, platelet derived growth factor; PDGFR, platelet derived growth factor receptor; TGF-β, transforming growth factor beta; RAR, retinoic acid receptor; TNF-α, tumor necrosis growth factor alpha.

Principles of Hair Follicle Morphogenesis

91

Table 6.3. Transgenic and knockout mice affecting hair follicle development Molecule

Method

Effect

Activin βA Activin βB BMP-4 EGFR FGF-7 (KGF) FGF-7 KGFR (dominant negative) Follastatin IGF-I/IGF-II IGF-I IL-6 PTHrp TGF-α TGF-α TGF-β1 α5/β1 integrin

Knockout Knockout Transgenic Knockout Knockout Transgenic Transgenic

Follicles fail to develop143 Follicles fail to develop143 Smaller follicles88 Curly hair and disorientation of follicles99 Abnormal hair fiber formation107 Suppression of follicle formation144 Decreased follicle number and abnormal morphology106 Abnormal follicle development145 Abnormal follicle development116 Premature follicle development117 Retards hair growth146 Disturbed hair follicle development147 Wavy hair fibers96 Stunted hair growth148 Reduction in number of follicles149 Hair follicles disorganized and stunted150

Knockout Knockout Transgenic Transgenic Transgenic Knockout Transgenic Transgenic Transgenic

Expression of Molecules During Human Hair Follicle Development A number of elegant studies have been carried out to characterize changes in patterns of extracellular matrix and cell adhesion molecules during embryonic development of human hair follicles.54,61 Prior to hair follicle development (60 days EGA) the embryonic skin is composed of a simple two layered epidermis composed of the basal cells and periderm separated from the dermis by a simple dermal-epidermal junction (DEJ). The dermis is composed of the reticular lamina, which forms a thin mesenchymal layer beneath the basement membrane (BM), and the compact layer. As the epidermis begins to stratify (60-70 days EGA), the dermal epidermal junction (DEJ) also undergoes marked changes to give rise to the structurally complex DEJ. Prior to development of the hair follicle the epidermis is organized into a basal, intermediate and periderm layer and the dermis is composed of a loose extracellular matrix (Fig. 6.1). During this early stage of development the basal epidermal cells express keratins K5 and K14 as well as the hyperproliferative keratin K17 and keratin K19, a possible marker for stem cells.45,46 The basal cells of the epidermis are also positive for the EGF receptor (EGFR). The DEJ, which forms a complex BM, is composed of heparan sulfate proteoglycan (HSPG), chondroitin 4- and 6-sulfated proteoglycans (C4-SPG, C6-SPG), Perlecan and type IV collagen. The DEJ is negative for tenascin.54 The reticular lamina, which forms a thin layer of mesenchyme beneath the BM, stains positive for types I, III, V, VI and VII collagen as well as fibrillin. The compact mesenchyme, which includes the reticular lamina, stains positive for C4-SPG, C6-SPG and Perlecan, and is reported to be rich in growth factor receptors and cell adhesion molecules including p75 (the low affinity neurotrophin receptor), neural cell adhesion receptor (NCAM), the PDGF-α and PDGF-β receptor subunits.61 In addition to these molecules, the compact mesenchyme of other animals has also been reported to express fibronectin, transforming growth factor β

92

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 6.3. Possible hypothesis for the roles of some growth factors and signaling molecules in the early stages of hair follicle development. We propose that early expression of the TGF-βII receptor within the epidermis prior to follicle development indicates the site of future placode development and subsequent follicle formation. However, follicle development will not take place unless a complex pattern of reciprocal signaling between the epidermis and dermis occurs. Candidate molecules within the epidermis may include members of the FGF family of growth factors including FGF-1, -2 and -4. FGF expression in the epidermis may then induce Shh expression which signals to both the epidermis and dermis. Candidate molecules that may be induced by Shh include TGF-β1, BMP-2 and BMP-4. TGF-β1 binds to the TGF-βII receptor and in so doing downregulates PKC, which in turn limits expansion of the interfollicular domain and which may favor appendage formation, thus establishing the boundary of the epithelial placode. TGF-β1 and transient expression of BMP-4 in the dermis stimulate mesenchymal aggregation and hence dermal papilla formation. Once the dermal papilla has formed it is then able to signal back to the epidermis and stimulate epithelial downgrowth. Reciprocal signaling of Lef1 between the dermal papilla and the placode appears to be important for the stabilization of the placode and maintenance of the dermal papilla. Subsequent downgrowth of the epithelial placode may be controlled by the production of a number of putative signaling molecules by the dermal papilla. Molecules produced by the dermal papilla and epithelial downgrowth at this time include FGF-7 (KG) and insulin-like growth factors (IGFs).

(TGF-β), acidic and basic fibroblast growth factor (aFGF and bFGF), bFGF receptor (bFGFR), mRNA for bone morphogenic protein 2a and 4 (BMP-2a, -4) and hyaluronic acid.61 Follicle initiation: Pregerm and hair germ formation Prior to formation of the hair germ, subtle changes take place in the BM, which suggest that it undergoes modification to support the downgrowth of the hair germ. These changes in the BM are characterized by the appearance of tenascin and a marked increase in expression of C6-SPG in regions which correlate with the sites above which epithelial placodes will form. The interfollicular epidermis remains negative for tenascin and expression of C6-SPG is also reported to decrease.54 Tenascin, a glycoprotein component of extracellular matrix, has been widely implicated in epithelial-mesenchymal interactions in a number of tissues. In the developing tooth, which undergoes epithelial-mesenchymal interactions analogous to those seen during hair follicle development, it has been shown that dental epithelium can stimulate tenascin synthesis within the mesenchyme.54

Principles of Hair Follicle Morphogenesis

93

Thus it would appear that prior to epithelial placode formation the mesenchyme beneath the subsequent site of development is modified in preparation for appendage formation. This data suggests that even prior to epithelial placode formation, the epithelium is able to direct the mesenchyme in preparation for appendage formation (see later). This would appear to correlate well with the development of the feather, in which prior to formation of the feather germ tenascin mRNA and protein are detected in epithelial cells of the follicle placode.62 With the onset of formation of the hair germ there is a marked decrease in EGF receptor (EGFR) expression in the developing epithelial placode. Whether this loss of (EGFR) expression is a result of downregulation of EGFR is not clear, as EGFR localization was determined by [125I] EGF ligand binding and it is possible that EGFR expression was masked by occupation of the EGFR by endogenous EGF.63 The mesenchymal cells that form the dermal aggregate beneath the epithelial placode stain strongly for NCAM during hair germ formation, whereas the interfollicular dermis stains only weakly. The extracellular matrix surrounding the mesenchymal aggregates is rich in NCAM, tenascin, p75, α and β subunits of the PDGFR, C6-SPG; but is negative for collagens type I, III and V. The epithelial placode that forms the hair germ stains weakly for E-cadherin in contrast with the suprabasal cells, which are strongly positive. ICAM staining is reported weakly on the outer cells of the hair germ. However, this expression is only transient and is lost at the hair peg stage. Subsequent follicle morphogenesis During the peg stage of hair follicle development, changes in the patterns of keratin expression occur. Keratins K5/K14 as well as K17 become restricted to the outer layers of the hair follicle peg, whereas keratin K19 becomes localized to restricted regions of the hair follicle, including the bulge.45 PDGF-α is expressed by the basal cells of the epidermis whereas the β chain (PDGF-β) is expressed by the supra basal cells. As the hair follicle peg stage develops, both PDGF-α and PDGF-β continue to be expressed, with the PDGF-α expression occurring in the outer epithelial cells of the developing follicle and PDGF-β in the inner cells. During early embryonic development of the epidermis, the basal cells and periderm express E-cadherin. During the peg stage of hair follicle development, E-cadherin is expressed in all epithelial components of the follicle apart from the epithelial cells that will give rise to the hair follicle matrix. In addition to lacking E-cadherin, the presumptive matrix cells are also negative for the basal keratins K14 and K5. ICAM staining is lost from the epithelial cells. The BM of the hair peg remains very strongly positive for tenascin, whilst staining in the interfollicular BM remains weak. During the formation of the hair peg, gradients of tenascin expression around the developing follicle have been reported.54 Staining for tenascin extends deeper from the follicular BM into the interfollicular dermis on the anterior side of the follicle than the posterior. In addition a proximal-distal gradient is also established in the follicle sheath, with strong labeling near the epidermis which becomes progressively weaker towards the dermal papilla, which only shows weak staining for tenascin.54 Staining for C6-SPG remains strong in the BM of the hair peg in contrast to the weaker staining in the interfollicular BM. Thus it would appear that C6-SPG may act as a useful marker to delineate between follicular and interfollicular BM. Both the follicular and interfollicular BM stain weakly for C4-SPG, as does the interfollicular dermis. The connective tissue sheath and the dermal papilla of the hair follicle are, however, negative for C4-SPG. In contrast to the apparent proximal-distal gradient of tenascin expression in the developing follicle, NCAM expression shows a reverse gradient; the dermal papilla has strong staining for NCAM, which then becomes subsequently weaker towards the epidermis.54

94

Molecular Basis of Epithelial Appendage Morphogenesis

During the bulbous peg stage of hair follicle development, germinative epithelial cells within the follicle bulb divide and give rise to daughter cells. These daughter cells form the matrix and undergo lineage restricted differentiation, giving rise to the component structures of the hair fiber (medulla, cortex and cuticle) and IRS (cuticle Huxley’s and Henle’s layers). Follicle sculpture by apoptosis As a recurrent theme in the embryogenesis of all organs, deletion of individual cells by programmed cell death (apoptosis) is also utilized as a tool for sculpturing the developing hair bulb.64 During the formation of the hair cone, transglutaminase I-positive cells and apoptotic cells identified by in situ end labeling (TUNEL stain) appear in this region.65 This suggests that during hair follicle development apoptosis may be restricted to the central core of epithelial cells that undergo terminal differentiation and/or are deleted during formation of the IRS and hair canal. These observations support the detailed EM studies of Holbrook and Odland,39 Robins and Breathnach44 and Hashimoto.49 Interestingly, we have recently also identified TUNEL+ cells in the central IRS of fully mature anagen hair follicles in mice.66 This observation begs us to ask the question Are developmental patterns of programmed cell death in the hair follicle carried over into the hair cycle? In the lanugo follicle, the anti-apoptotic mitochondrial protooncogene product, Bcl-2,67 is restricted to the epithelial cells within the bulge,65 though epithelial Bcl-2 expression is clearly not a stem cell marker, since it is also seen in non-stem cell regions of the murine follicle epithelium.66,68 However, it is interesting to note that Bcl-2 is strongly expressed in the developing epithelial placode, while only limited, sporadic Bcl-2 expression is seen in the interfollicular human epidermis.65 As the hair germ forms, Bcl-2 becomes expressed in the “invaginating” epidermis, the presumptive matrix and the mesenchymal dermal papilla. By the hair peg stage, Bcl-2 is restricted to the dermal papilla and matrix cells adjacent to the dermal papilla.65 Though we still know pitifully little on the exact choreography, relative importance and control of apoptosis during follicle morphogenesis, this may be one of the basic forces controlling hair follicle morphogenesis. We speculate that many congenital hair growth abnormalities in essence reflect abnormalities in follicular apoptosis during development, and many of the signaling molecules implicated in the control of hair follicle morphogenesis (Table 6.2)may exert at least some of their developmental effects via the modulation of apoptosis.19,66

Molecular Controls of Murine Hair Follicle Development Multiple spontaneously arisen or experimentally generated mouse strains with well-defined genetic abnormalities also show abnormalities in the development of all or selected hair follicles (Table 6.3 and see refs. 32, 33). Yet, as will become evident below, even the availability of such invaluable hair research tools has still not stimulated the proposal of a unifying hypothesis that comprehensively explains how hair follicle induction and morphogenesis might operate on the molecular level. Part of the problem is that these hair phenotypes of mouse mutants were commonly generated by immunologists, developmental, neuro- and molecular biologists without a primary interest in hair, and thus were not subjected to rigorous trichological analyses. Consequently, the hair phenotypes of these mice are frequently reported only as a mere footnote. In order to make optimal hair research use of these intriguing mutants, we advocate subjecting all mouse mutants—irrespective of whether or not they display an obvious hair phenotype—to a systemic screening for defined, quantifiable differences in the time course and morphology of neonatal hair follicle morphogenesis (comparing mutant and wildtype

Principles of Hair Follicle Morphogenesis

95

littermates). This is most conveniently done by using a simple, pragmatic classification scheme, and should greatly aid in the clarification of the specific developmental role(s) of any of the genes and gene products discussed below. Homeobox (Hox) genes and homeoproteins Homeobox genes are essential for the patterning and development of many segmental structures in both invertebrates and vertebrates.5 Because Hox genes play such a crucial role in pattern formation,5 Chuong et al69 suggested that Hox genes may also play an important role in the phenotypic determination of skin appendages. He went on to show in the avian system that asynchronous expression of a macrogradient of different Hox genes can lead to a large repertoire of different expressions and hence different feather types (see chapter 5). In murine skin, the patterns of homeobox gene expression were found to vary both temporally and spatially within the developing skin and hair follicles.70 In embryonic mouse skin, HoxC8 was expressed according to two graded patterns along the cephalo-caudal and dorsal-ventral axis in both the epithelial cells of the developing hair follicle and in the dermal papilla. Hox d9 and d11 were expressed during the formation of the hair bud, where their expression was restricted to the epithelial cells of the hair follicle. Both Hox d9 and d11 remained expressed in the fully developed hair, where they were restricted to the differentiating matrix cells of the hair follicle bulb. Hox d11 was also strongly expressed in the basal cells of the ORS. Expression of both Hox d9 and d11 were switched off with the onset of catagen. Unfortunately, the authors did not investigate whether expression was switched on again with the onset of the development of the new anagen hair follicle. In contrast to expression of Hox d9 and d11, Hox d13 was expressed relatively late in hair follicle development and was restricted to the differentiating matrix cells of the follicle bulb. Moreover, once the hair follicle was fully developed Hox d13 expression was switched off.70 These patterns of HoxD expression suggest that they may play an important role in specifying the timing, location and duration of hair follicle development. It is especially intriguing that both Hox d9 and Hox d11 were expressed in the fully developed follicle, but were switched off with the onset of catagen. Perhaps, these Hox genes play an important role in the timing and duration of the active, anagen, stage of the hair growth cycle. Other Hox genes that appear in the initial stages of hair follicle development include Msx-1 and Msx-2.71,72 In tooth development, Msx-1 is expressed early in the dental mesenchyme and is believed to play an important role, along with Lef1 and activin, in signaling formation of the enamel knot (see chapters 9, 14). In the feather and hair follicle, Msx-1 and Msx-2 are expressed in the epithelial placode.73 Both Msx-1 (Hox-7) and Msx-2 (Hox-8) reportedly are expressed in the developing whisker follicle.71,72 However, their exact follicular expression patterns were not described, making it difficult to judge their role in follicle development. Therefore one can currently only be guided by drawing analogies to feather development (chapters 4,5). Here, the functional importance of Msx gene expression in placode formation has been demonstrated by experiments showing that the culture of embryonal chicken skin in the presence of either forskolin, an activator of adenyl cyclase, or dibutyryl cyclic adenosine monophosphate, an activator of PKA, inhibits feather bud elongation, associated with a marked decrease in Msx-1 and Msx-2 expression.73 Interestingly, in the developing feather bud, Msx gene expression73 co-localizes with that of tenascin (see above). Retinoic acid Retinoic acid (RA) is recognized as an important modulator of embryogenesis and has marked homeotic effects on skin appendage formation.4,74,75

96

Molecular Basis of Epithelial Appendage Morphogenesis

When embryonic lip skin from the mouse is maintained with exogenous RA, instead of vibrissae follicle development skin appendages with a glandular structure were observed.74 However, the effects of RA on skin are only observed if skin is treated early in the initiation stage, during the formation of the epithelial placode and aggregation of the mesenchyme. Tissue recombinant experiments in which embryonic mouse lip skin, separated into dermis and epidermis, and untreated epidermis placed on treated dermis, was cultured with retinoids suggest that retinoids are acting via the dermis to initiate a switch from follicular to glandular morphogenesis.75 Three nuclear receptors for RA have been identified (RARα, RARβ and RARγ). RARα is expressed in the developing dermal papilla of 12.5 day old mice whilst RARγ is expressed in both the papilla and the epithelial placode. During subsequent follicle development, RARα and RARγ expression were switched off in the dermal papilla, but the epithelial component of the follicle continued to express RARγ.76 These observations suggest that RA plays an important role in skin appendage morphogenesis and subsequent differentiation of follicle keratinocytes. This is supported by the observation that RARβ expression appears to be restricted to glandular morphogenesis in nasal tissue and that in RA treated skin, which is capable of stimulating glandular morphogenesis in embryonic follicles, there is a marked increase in RARβ gene expression.4 In addition to the RA nuclear receptors, a cytoplasmic RA-binding protein (CRABP) has been identified in the connective tissue sheath surrounding the hair follicle.77 The role of this CRABP in the hair follicle is unknown, but it may be involved in sequestering free RA.4 Transforming growth factor beta (TGF-β) and bone morphogenic proteins (BMP) Members of the transforming growth factor TGF-β/BMP superfamily play important roles in many developmental processes.5,78,79 In the skin, TGF-β isoforms are now acknowledged as key modulators of keratinocyte proliferation, differentiation and apoptosis, of fibroblast proliferation and extracellular matrix remodeling, and as angiogenic factors.8,80 Immunohistochemistry has localized TGF-β1, TGF-β2 and TGF-β3 to the developing hair follicle of mice.81 However these studies did not identify precisely which hair follicle compartments were positive for these growth factors and whether there were any spatial or temporal expression differences. During hair follicle development in the mouse TGF-β2 mRNA is first expressed in the dermis (15.5 dpc), but by day 18.5 dpc mRNA expression is switched from the dermis to the epidermis.81 In contrast, TGF-β2 protein is expressed in both the dermis and epidermis of 15.5 dpc mice.81 This suggests that TGF-β2 protein produced by the dermis may be able to bind to receptors both in the dermis and epidermis. This is interesting, as the type II receptor for TGF-β is expressed very early in the epidermis prior to placode visible formation (see Fig. 6.2). In addition, the switch in TGF-β2 mRNA expression between the dermis and epidermis also appears to mimic the switching seen during the development of other skin derived appendages, although whether this occurs in the hair follicle has yet to be established. Moreover, it must also be remembered that these observations are based on two different sets of experiments and these patterns of expression remain to be established using identical tissues. Recently, we have established by immunohistology the expression patterns of both major TGF-β receptors.23 The obtained immunoreactivity patterns suggest that, in murine skin only follicle keratinocytes, and not epidermal keratinocytes, sebocytes, or extra-epithelial cells, make substantial usage of signaling mediated by the TGF-β-RI/TGF-βRII complex, and that ORS keratinocytes and a selected hair matrix cell population are the main targets of TGF-β1 and TGF-β3 signaling.23 Interestingly, type II TGF-β receptor proteins are selectively expressed in those sectors of the epidermis destined to form a hair follicle, even before the placode formation is mor-

Principles of Hair Follicle Morphogenesis

97

phologically recognizable. This makes TGF-β type II receptor an excellent light microscopy marker for imminent follicle development—and the first growth factor receptor recognized so far that allows one to identify an area of epidermis that will later on form a hair placode, while the area is morphologically still indistinguishable from the neighboring epidermis.23 Unfortunately, it is not yet possible to say with any certainty from which tissue compartment the corresponding high affinity ligands for this receptor (TGF-β1, -3) arise during follicle morphogenesis, and what this means for the dispute about where the key signals of hair follicle induction originate: while a mesenchymal origin of TGF-β1, and/or -3 would be in line with the concept that the dermis sends the first message (“make an appendage”) to the appropriately receptive (i.e., TGF-β receptor type II-expressing) epithelium, it remains open which signals control this strikingly selective epithelial receptor expression (see below, Lef1). Disruption of TGF-β1gene expression in mice apparently has no gross effect on development However, after two to three weeks these mice die from a rapid wasting syndrome.83,84 Because TGF-β has been strongly implicated in many developmental processes, the authors have proposed that the failure of TGF-β null mice to show any gross developmental abnormalities may be due to maternal transfer of TGF-β across the placenta. Transgenic mice that overexpress TGF-β in the epidermis produce a thin epidermis and fewer hair follicles. This is not surprising, as TGF-β is a potent inhibitor of epithelial proliferation and has been shown to inhibit hair follicle growth in vitro.85 The BMPs are expressed during embryonic development of nearly all mammalian systems and are believed to play important roles in both the regulation of cell proliferation and terminal differentiation.79 It has been shown by in situ hybridization and immunohistochemistry that the spatio-temporal expression of BMP-2 and BMP-4 correlate with early hair follicle, tooth and feather induction. Reportedly, BMP-4 is expressed in the mesenchyme around day 12 of gestation, prior to epidermal placode formation; however, expression is transient and no signal is detected following downgrowth of the epidermal bud.86 In the developing mouse whisker follicle, BMP-2 is localized to the epidermal placode at day 13.5 gestation, and as the follicle develops expression remains localized to the hair matrix cells and the hair cone of the IRS.33 BMP-4 is also expressed in the developing hair follicle, although its pattern of expression is transient and restricted to the mesenchymal condensation underlying the epidermal placode;86 this contrasts with BMP-2. During subsequent follicle development, expression of BMP-4 is lost. The transient nature of BMP-4 expression is unclear; however, similar but more detailed patterns of BMP have also been reported in the developing tooth87 (see chapter 14). This suggests that in the tooth BMP-4 expression in the epidermis signals placode formation and also switches on BMP-4 expression in the underlying mesenchyme. Moreover, agarose beads soaked in BMP-4 are able to induce BMP-4, Msx-1 and Msx-2 gene expression in isolated dental mesenchyme.87 Since Msx-1 and Msx-2 are some of the earliest genes transcribed during tooth and hair follicle development71,72 this suggests that during hair follicle development, BMP expression in the epidermis may be an important inducer of BMP-4 and Msx genes in the hair follicle mesenchyme. In transgenic mice where BMP-4 has been ectopically expressed in the ORS of pelage and vibrissae follicles using the promoter region of bovine keratin IV, abnormal hair follicle growth is reported.88 In these transgenic follicles cell proliferation in both the ORS and matrix cells is inhibited, and this appears to result in a failure of hair follicles to initiate a second hair growth cycle, resulting in alopecia. In BMP-4 transgenic mice early hair follicle development appears to occur normally, and follicle defects are only observed as follicle cells begin to differentiate into ORS.88 However, these BMP-4 transgenic mice are difficult to interpret. BMP-4 is normally transiently expressed in the mesenchyme of the developing

98

Molecular Basis of Epithelial Appendage Morphogenesis

hair follicle and not in the ORS. Therefore, ectopic expression of BMP-4 in the ORS is likely to install abnormal signaling patterns within the follicle that may well markedly disrupt normal patterns of hair follicle growth and cycling. Both BMP-2 and BMP-4 are reported to have similar effects on cartilage and bone formation, and Blessing et al88 have suggested that in their transgenic mice BMP-4 may substitute for, and exert a similar biological activity to, BMP-2. BMP-2 has been proposed as an inhibitor of cell proliferation. Further, the observation that ectopic expression of BMP-4 in the ORS inhibits cell proliferation in fully developed hair follicles, but apparently has no effect on proliferation during hair follicle development during which both BMP-2 and 4 are expressed, suggests that tissue specific expression of Type I and Type II BMP receptors may play an important role in regulating the activity of these molecules both spacially and temporally within the hair follicle. In addition to BMP-2 and -4, two other BMP have also been detected in developing hair follicles.89 BMP-3 was weakly expressed in the dermal papilla and in the ORS and IRS at day E15. By day E18, expression was only detected in the IRS. BMP-7 mRNA was absent from the hair follicle at day E15, but expression was detected in the ORS and IRS at E18; in addition weak expression was also observed in the connective tissue sheath. During the later stages of hair follicle development (5 days postnatal) BMP-7 mRNA was observed in both the dermal papilla and in the lower ORS.89 Control of BMP expression: Hedgehog and patched Upstream regulation of BMPs is poorly understood in vertebrates. However, there is evidence that the BMPs are regulated by members of the hedgehog family. In Drosophila, Dpp is regulated by hedgehog (hh) and several other components of this pathway have been identified, including patched (ptc) which encodes a transmembrane protein and cubitus interuptus (ci), a zinc finger transcription factor. To date, however, the exact details of how these components interact is not known. Currently three mammalian homologues of the hedgehog have been identified, Sonic hedgehog (Shh), Desert hedgehog (Dhh) and Indian hedgehog (Ihh). In chick limb buds ectopic expression of Shh induces expression of HoxD genes and BMPs. Moreover, ectopic expression of Shh also upregulates expression of patched (ptc), which suggests that a similar signaling pathway between Shh and BMPs may also exist in vertebrates (see below). This is further supported by the recent cloning of three mammalian genes related to Drosophila ci (Gli, Gli2 and Gli3). Whether these genes play a role in regulating BMPs is still to be established, and their role, if any, in hair follicle development is not known. The action of Shh appears to be mediated by opposing the repressive activity of the gene patched (ptc), which codes for a protein with multiple transmembrane domains. However, as hh induces ptc, an interesting negative autoregulation loop appears to operate. In the developing whisker, both Shh and ptc are expressed in complementary sites, with Shh expression being surrounded by a ring of ptc expression. This ptc would appear to form an outer boundary that restricts Shh activity.90 Retroviral ectopic expression of Shh in chick skin results in the formation of extra-large feather buds with aberrant or lost polarity.91 This data suggests that Shh and ptc work together to control the size of the developing skin appendage and this may be an important determinant of follicle size. Regulation of Shh expression in the epidermis appears to be under the control of the mesenchyme. Whether the mesenchyme is first conditioned by the epidermis is still open to debate. Ting-Bereth and Chuong91 (and see chapter 13) have proposed that as yet unidentified molecules in the mesenchyme signal Shh expression in the epithelial placodes. Shh protein expressed in the epithelium then induces dermal condensation (presumptive DP)

Principles of Hair Follicle Morphogenesis

99

mediated via TGF-β2. Whether this pathway also operates during hair follicle development remains to be determined. Epidermal growth factor and transforming growth factor α EGF receptor expression, as determined by the binding of exogenous [125I]EGF, has been demonstrated in both embryonic human92 and murine skin.63 In both human and murine skin, EGF receptors appear to be present on the basal cells of the epidermis prior to formation of the epithelial placode. However, as hair follicle development occurs, EGF receptors are lost from the basal epithelial cells which give rise to the epithelial placode and under which mesenchymal cells aggregate.63 It is not known, however, whether this loss of EGF receptors resembles downregulation of receptor expression or occupancy by endogenous EGF. During subsequent downgrowth and development of the hair germ intense EGF binding is detected on the epithelial component of the hair germ. During subsequent development of the hair follicle, EGF receptors are localized to the ORS and sebaceous gland of the hair follicle. The role of EGF in hair follicle development is still open to debate. Receptor expression would suggest that it plays an important role in proliferation of the epithelial cells of the hair germ that give rise to the anagen hair follicle. EGF has been shown to be highly mitogenic for ORS keratinocytes in cultured human hair follicles,93 and injection of EGF into newborn mice inhibits hair follicle development.94 Although the mechanism of this inhibition is not known, high doses of exogenous EGF may disrupt normal patterns of proliferation in the developing follicle in a similar fashion to that seen in adult follicles in vivo94 and in human hair follicles in vitro.93 It is still unclear whether EGF is the active ligand during hair follicle development, as EGF synthesis in mice is not believed to start until two weeks post partum. In contrast, TGF-α is synthesized earlier and appears to have identical effects to EGF (see ref. 25). TGF-α knockout mice show pronounced waviness of the hair fibers produced by both pelage and vibrissae follicles. Moreover, the hair phenotype of TGF-a knockout mice is identical to that seen in the spontaneous mouse mutation waved-1.96 Furthermore, the waved-2 mutant, which also produces a wavy coat, has been identified as a point mutation within the kinase domain of the EGF receptor.97 As a result of this mutation, EGFR signaling is compromised.98 Data from these mouse mutants suggests that EGF/TGF-α play an important role in positioning of the hair follicle within the skin (see below). This is confirmed by the data of Threadgill et al,99 who have shown that null mutation of the mouse EGF receptor (Egfr) results in the production of hair follicles which appear normal at birth but which, by day P5 after birth, appear irregularly placed and grow in a disorganized manner. Histology of hair follicles from these mice show premature separation of hair fiber from the IRS and premature keratinization of the IRS and hair fiber. Staining for keratin K6 shows abnormal expression in the precortical cells of the follicle bulb, and patterns of integrin staining showed abnormal expression of α3β1 and α6β4 integrins along the ORS of hair follicles. This suggests that EGFR ligands may regulate, via integrins, hair follicle development and positioning, and highlights the importance of topobiology for hair follicle morphogenesis (see chapter 15).99 Though EGF/TGF-α appears to play an important role in positioning of the hair follicle within the skin, its exact role in hair follicle development is still unclear. The Tabby (Ta) mutation in mice results in a reduction in the number of vibrissae follicle and teeth, abnormal coat texture and an absence of sweat glands. Gruneberg100 has shown that the Ta mutation appears to affect structures that arise as a result of epithelial mesenchymal interactions that give rise to epidermal downgrowth. Moreover, tissue recombinant studies101 have shown

100

Molecular Basis of Epithelial Appendage Morphogenesis

that the Ta defect resides within the epidermis, and that dermis from Ta mice is able to support hair follicle development. Histology of hair follicle development in Tabby mice shows that follicle initiation begins three days later than controls and finishes two days earlier. Furthermore it appears that only primary follicles are established and that these follicles produce only a hair fiber described as abnormal awl.101,102 The site of defect in Ta mice is not known. However, Kapalanga and Blecher103 have shown that some effects of the Ta mutation, such as delayed eyelid opening and incisor eruption, are reversed by exogenous EGF.103 This data suggests that the Ta gene may play a regulatory role in the EGF/TGF-α pathway. In addition, Blecher et al104 have shown that exogenous EGF will also induce sweat gland development in murine X-linked anhydrotic ectodermal dysplasia.104 Fibroblast growth factors Other early signaling molecules in the epithelial placode include members of the FGF family. Widelitz et al105 have shown that when small porous beads are soaked in FGFs and implanted into chick skin, new feather buds are induced. Moreover, these new buds express Shh as well as intracellular signaling molecules such as Grb, Ras, Raf, Erk.105 These molecules are usually expressed downstream of tyrosine kinase receptors such as the FGF receptor, and suggest that the FGFs play an important role in the formation of skin appendage domains.105 FGF signaling has also been implicated as an early event in limb formation, with FGF-4 inducing Shh and Msx in developing limb buds (see chapter 2). Transgenic mice which express a dominant negative KGFR targeted to basal and ORS keratinocytes resulted in a decrease in hair follicle numbers and abnormal hair follicle morphology.106 In contrast, KGF null mutant mice are reported to resemble the rough (ro) mutant mouse107 and are characterized by normal coats which progressively become matted. Histology of hair fibers, however, appears normal. Recombinant KGF has been shown to induce a dose dependent hair growth over the entire body of athymic nude mice (nu/nu). Histology of KGF treated mice showed increased proliferation of hair follicle matrix cells and a reversal of the nu/nu keratinization defect.108 In the developing hair follicle, two FGF receptor genes have been studied using in situ hybridization.109 FGFR1 mRNA was detected in the dermal papilla of 16.5 dpc hair follicles, whereas the FGF-2 receptor mRNA was expressed both in the dermal papilla and in the epithelial cells of the follicle bulb.109 This pattern of expression contrasts with that reported by Rosenquist and Martin110 for the fully developed anagen hair follicle, in which mRNA for the FGFR1 was expressed in the dermal papilla but the FGFR2 was only expressed in the matrix cells and not the dermal papilla. FGF-1 (aFGF) has been detected in the epithelial placode.111,112 FGF-2 (bFGF) is expressed in the dermal papilla at the follicle peg stage of development. FGF-7 (KGF) was found in the connective tissue sheath surrounding the follicle, but not in the dermal papilla, whereas the KGF receptor (KGFR) was detected in the hair follicle epithelium.113 However, in the fully developed anagen follicle, KGF mRNA was detected in the dermal papilla and KGF receptor mRNA is expressed in the hair follicle matrix cells.113 FGF-4 plays an important role in development but remains to be studied during hair follicle morphogenesis. However, in fully developed anagen hair follicles FGF-4 does not seem to be expressed.110 FGF-4 has, however, been strongly implicated in epithelial-mesenchymal interactions during development of many organ systems, but especially within the developing limb bud, tooth114 and feather.105 It is likely, therefore, that FGF-4 may well play an important role in development of the hair follicle and this remains to be studied.

Principles of Hair Follicle Morphogenesis

101

Insulin-like growth factors The insulin like growth factors IGF-I and IGF-II also play an important role in embryonic development.115 Mice carrying null mutations of the gene encoding the type I IGF receptor (IGF-Ir) have abnormal hair follicle development characterized by a marked decrease in the number of hair follicles.116 Moreover, hair follicles that were present in these mice were smaller and more widely spaced than wild-type controls. In contrast, mice carrying null mutations in the IGF-I gene have normal skin and hair follicle development. Likewise, IGF-II knockout mice also have normal skin development. However, double mutants containing knockout of both the Igf-I and IGF-II genes have abnormal skin and follicle development. It would thus appear that in the developing skin and hair follicle IGF-I and IGF-II are able to compensate and that they both act via the type I IGFR. The importance of IGF-I in hair follicle development is further demonstrated in transgenic mice in which overexpression of IGF-I cDNA was targeted to the skin using a human keratin 1 (K1) promoter construct (HK1). In mice overexpressing IGF-I, changes in the skin were characterized by epidermal hyperplasia and hyperkeratosis as well as premature hair follicle generation.117 Thus IGF-1 may be an important regulator of follicle development following hairgerm formation. Lef1 and other transcription factors Lymphoid enhancer-binding factor (Lef1) is a cell type-specific transcription factor expressed in lymphocytes of mice as well as in the neural crest derived mesenchyme, including that of tooth and whisker follicles. Lef1 gene expression has been detected in both the epithelium and mesenchyme during appendage formation.118,119 In the developing tooth Lef1 is first expressed in the epithelial placode (E10 and 11), but then shifts to the underlying mesenchyme (E12). During subsequent development of the tooth, Lef1 continues to be expressed in both the mesenchyme and epithelium. Lef1 may not, however, be essential for epithelial placode formation during tooth development, as Lef1-/- mice initiate the formation of tooth germs, although these germs do not develop beyond the bud stage due to failure of the DP to form. Thus, in the tooth, formation of the epithelial primordium is not dependent upon Lef1 gene expression. However, development of the tooth mesenchymal papilla and subsequent appendage development are dependent upon transient expression of Lef1 gene expression in the epithelium. In the developing whisker follicle, Lef1 protein expression appears first in the mesenchyme (E11); whether Lef1 gene expression occurs in the epithelium at this stage has not been reported. However, this pattern of expression would appear to contrast with the tooth, in which transcripts of Lef1 are first detected in the epithelial placode. However, following initiation of placode formation Lef1 is also detected in the epithelium and remains expressed in both the epithelium and mesenchyme during the subsequent stages of follicle development. However, in contrast to tooth development in Lef1-/- mice, which is arrested at the tooth germ stage, whisker development fails to be initiated. Thus, development of the whisker hair germ appears to be dependent upon Lef1. Moreover, recombinant studies have shown that development of the hair germ is dependent upon mesenchymal Lef1 expression between E11 and E12. Finally, Lef1 gene expression can be activated by BMP-4. Thus early signalling events in hair placcode formation may involve shh→BMP→Lef1. Whn The product of the mouse nude locus Whn is the mutated gene responsible for the nude mouse and nude rat phenotype.32 The winged helix nude (Whn) gene codes for a transcription factor belonging to the to the winged helix domain that can suppress expression of differentiation responsive genes, and as such regulates the balance between epithelial

102

Molecular Basis of Epithelial Appendage Morphogenesis

cell growth and differentiation.120 In hair follicles Whn has been detected in the epithelial cells.120 Similarly, the long elusive product of the murine hairless gene, whose functional deletion leads to progressive alopecia and follicle loss in the third week of postnatal life,32,121 is now thought to be a transcription factor of the zinc finger protein family which is expressed in the epidermis and follicle of normal mice.34 Here, it may be an essential factor in maintaining dermal papilla and ORS integrity, and in controlling intrafollicular apoptosis.122 M-twist M-Twist is a basic helix loop helix (bHLH) transcription factor that is believed to play an important role in inhibiting muscle development and chondrogenesis. However, its pattern of expression also suggests that it may play an important role in epithelial-mesenchymal interactions. In the mouse, M-Twist is expressed in the mesenchyme of the developing tooth and in the developing whisker follicle. Its role in the development of these tissues is not clear; however, mesenchyme of both the DP of the tooth and whisker follicle is derived from the cranial neural crest, and it has been suggested that M-Twist may play a role in regulating the differentiation of the DP.123 Id1, Id2, Id3 and Id4 The murine dominant negative helix-loop-helix (dnHLH) family of proteins, Id1, Id2, Id3 and Id4, are proteins that inhibit the activities of basic helix-loop-helix (bHLH) transcription factors. In developing whisker follicles124 Id1, Id2 and Id3 were detected in the condensing mesenchyme beneath the epidermal placode. Id4 was not detected during early hair follicle development. During subsequent follicle morphogenesis and cytodifferentiation (14.5-16.5 dpc) all three transcription factors were expressed in the ORS, connective tissue sheath and dermal papilla. Id4 was detected in the IRS.124 Notch Notch, a transmembrane signaling receptor first identified in Drosophila, and midkine (MK), a novel heparin binding growth factor encoded by a retinoic acid responsive gene, are expressed in undifferentiated epithelial and mesenchymal cells125,126 and play important roles in determining cell fate. Both Notch and midkine have been proposed as candidate competence factors. That is, they act by blocking cell differentiation, which maintains the competence of the undifferentiated cells in both the developing tooth and hair follicle to respond to inductive signals that determine their developmental fate.127,128 During embryogenesis Notch transcripts are expressed in tissues undergoing epithelial-mesenchymal interactions. In the hair follicle Notch is expressed in the hair bud.127 Notch and midkine are expressed in undifferentiated epithelial and mesenchymal cells. In the hair follicle Notch is expressed in the matrix cells of the hair follicle bulb, and not in the dermal papilla or connective tissue sheath. Moreover, expression in the matrix was restricted to cells that form the precursors of the hair fiber and IRS. Notch was not detected in the bulge or the ORS. These data support the hypothesis that Notch may be a competence factor, as its pattern of expression in the hair follicle appears to be restricted to cells that have left the proliferative pool but have yet to terminally differentiate.127,128 Serrate-1 (Jagged-1) Mouse Serrate-1 (jagged-1), a mammalian homologue of the Drosophila serrate-1 gene is believed to act as a ligand for the Notch receptor. In the developing whisker follicle Ser-1 transcripts are found first in the epithelium at E12.5 and then from E13.5-14.5 are found in both the epithelium and mesenchyme of the hair follicle.126 This pattern of expression is similar to that seen in the developing tooth, where expression is first detected in the dental

Principles of Hair Follicle Morphogenesis

103

epithelium at day E10.5-11 and later (E12-12.5); as expression in the epithelium is downregulated increased expression is observed in the mesenchyme. In the developing tooth Ser-1 expression in the dental mesenchyme can be induced by FGF-4 but not by BMP. This contrasts with Notch, which is induced in the mesenchyme by BMP but not FGF-4.126 Thus, in the developing tooth, receptors and ligands for the Notch pathway are induced by separate signaling pathways. Midkine and pleiotrophin Midkine and pleiotrophin are heparin binding growth factors associated with developing organs undergoing epithelial-mesenchymal interactions.125 Prior to hair follicle development, both midkine and pleiotrophin are expressed in the epidermis and on mesenchymal cells situated at the site of follicle induction. However, as hair follicle development takes place, midkine remains expressed in both the epithelium of the developing follicle and in the mesenchyme, whereas pleiotrophin expression is restricted to the epithelium. By 16.5 dpc midkine is restricted to a weak stain in the dermal papilla. Likewise pleiotrophin is only weakly expressed in the follicular epithelium and in occasional mesenchymal cells of the dermal papilla.125 Thus, both midkine and pleiotrophin appear to be expressed only during the follicular induction and development, and expression is much reduced during the later stages of morphogenesis and cytodifferentiation. Patterns of midkine and pleiotrophin expression appear to correlate with that of syndecan-1, which may serve to regulate the biological activity of these two molecules. Currently the role of midkine and pleiotrophin in hair follicle development is not known, although it has been suggested that they may be mitogens.125

A Working Model: Molecular Controls of Hair Follicle Morphogenesis The process of hair follicle embryogenesis may be considered to be composed of three developmental stages: 1. Induction; 2. Organgenesis; 3. Cytodifferentiation. It is clear that hair follicle development involves a series of tightly regulated reciprocal interactions between the epithelium and mesenchyme both prior to follicle induction and during subsequent morphogenesis. During the early stages of induction the reciprocal pattern of signaling would appear to be as follows: 1. Initial signal to make an appendage. (Source unclear: epithelium or mesenchyme?) 2. Epithelium signals to mesenchyme to prepare to make an appendage. 3. Mesenchyme signals back to the epithelium to form a hair placode. 4. Epithelium begins to form a placode and signals the mesenchyme to aggregate. 5. The mesenchyme begins to form a mesenchymal aggregate and signals back to the placode. 6. Hair germ development proceeds and signals to the mesenchyme to make a dermal papilla. 7. Subsequent morphogenesis of the hair follicle may now chiefly reside with the dermal papilla. Although the molecular mechanisms of hair follicle induction have yet to be established, there is a sufficient body of data, both from studies on the hair follicle and also from other skin appendages such as the tooth and feather, to permit us to propose a tentative pathway of signaling that may result in hair follicle induction and subsequent follicle morphogenesis and cytodifferentiation (Fig. 6.3).

104

Molecular Basis of Epithelial Appendage Morphogenesis

Although it is generally considered that the initial signal for hair follicle induction arises in the mesoderm, this has still to be established. In our model, we propose that an as yet unidentified early signal from either the mesoderm or ectoderm results in induction of members of the FGF family. FGF signaling then induces Shh, which is able to signal to both the ectoderm and the mesenchyme. In the mesenchyme, Shh is responsible for the transient expression of BMP-4 which may trigger the subsequent formation of the mesenchymal aggregate that will eventually give rise to the dermal papilla. The response of the mesenchyme to Shh signaling from the epithelium may well be dependent upon other permissive signals from the epithelium. Moreover, in addition to the transient expression of BMP-4, Shh may also induce TGF-β, Msx genes, Lef1 and other ‘architectural’ transcription factors within the mesenchyme that coordinate the expression/repression of genes required for normal hair follicle organogenesis (Fig. 6.3). Formation of the epithelial placode appears to be dependent upon Lef1 expression in the mesenchyme. Lef1 expression is reported to occur first in the mesenchyme, and is followed a day later in the placode. Therefore, Lef1 or transcription factors that are induced downstream of Lef1 may serve as permissive factors that inform the epithelium that dermal papilla formation has begun and that epithelial placode formation may proceed. During formation of the epithelial placode, Shh induces expression of BMP-2 and other downstream signaling molecules such as the HoxD genes, Msx genes and Lef1. During placode formation, Lef1, and probably other transcription factors produced by the placode, then signal to the mesenchymal aggregate to continue to form a dermal papilla. The dermal papilla may then take control of subsequent follicle morphogenesis and cytodifferentiation for example by the secretion of powerful morphogens such as TGF-β, HGF/SF and IGF-1. In addition to the simple scheme outlined here (Fig. 6.3), it is also clear that other important signaling pathways operate and play a major role in follicle formation. Early expression of TGF-βII receptor in the epithelium prior to placode formation, and before formation of the mesenchymal aggregate, suggests an important role for TGF-β. This role may be twofold. First, TGF-β is a potent inhibitor of epithelial cell proliferation and may be closely involved in the regulation of hair germ development. Second, in chick skin TGF-β is a potent inhibitor of PKA, and PKA expression has been associated with expansion of the interfollicular epithelium. The role of TGF-β in hair follicle development may therefore be to maintain the competence of the epithelium to respond to inductive signals from the mesenchyme and form a skin appendage. Finally, IGF-I has been strongly implicated in hair follicle development and, moreover, during limb development IGF-I has been shown to induce thickening of the epithelium, forming a structure similar to the apical ectodermal ridge (AER).151 Furthermore, it has also been reported that the ability of FGFs to stimulate outgrowth of the limb mesoderm is dependent upon IGF-I.152 It is therefore possible that during hair follicle induction IGF-I may interact with FGFs to promote epithelial placode formation and may also play an important role in subsequent mesenchymal aggregation.

References 1. Maderson PFA. Some speculation on the evolution of the vertebrate integument. Amer Zool 1971; 12:159-171. 2. Montagna W, Parakkal PF. The Structure and Function of Skin. New York: Academic Press, 1974. 3. Ebling FJG. Comparative dermatology. In: Champion RH. Textbook of Dermatology. Oxford: Blackwell, 1991. 4. Hardy MH. The secret life of the hair follicle. TIG 1992; 8:55-61.

Principles of Hair Follicle Morphogenesis

105

5. Gilbert SF. Developmental Biology. Sunderland, MA: Sinauer, 1994. 6. Spearman RIC. The mammalian epidermis and its derivatives. The evolution of mammalian keratinized structures. Symp Zool Soc London 1964, 12:67-81. 7. Moll I, Paus R, Moll R. Merkel cells in mouse skin: Intermediate filament pattern, localization, and hair cycle-dependent density. J Invest Dermatol 1996; 106:281-286. 8. Paus R, Peters EMJ, Eichmüller S et al. Neural mechanisms of hair growth control. J Invest Dermatol Sympos Proc 1997; 2:61-68. 9. Botchkarev VA, Eichmüller S, Johansson O et al. Hair cycle-dependent plasticity of skin and hair follicle innervation in normal murine skin. J Comp Neurol 1997; 386:379-395. 10. Koelliker A. Zur Entwicklungsgeschichte der ausseren Haut. Z wiss Zool 1850; 2:67-92. 11. Unna PG. Beitrage zur Histologie und Entwicklungsgeschichte der menschlichen Oberhaut und ihrer Anhangsgebilde. Arch mikroskop Anat Entwicklungsmechan 1876; 12:665-741. 12. Stohr P. Entwicklungsgeschichte des menschlichen Wollhaares. Anat Hefte Abt 1903; 123:1-66. 13. Dry FW. The coat of the mouse (Mus musculus). J Genet 1926; 16:287-340. 14. Pinkus F. The development of the integument. In: Keibel H, Mall F, eds. Manual of Human Embryology. Vol I. Philadelphia: Lippincott, 1910:243-291. 15. Pinkus F. Die normale Anatomie der Haut. In: Jadassohns Handbuch der Haut- und Geschlechtskrankheiten. Berlin: Springer, Vol. 1/I, pp. 1-378, 1927. 16. Pinkus H. Embryology of hair. In: Montagna W, Ellis RA, eds. The Biology of Hair Growth. New York: Academic Press, 1958:1-32. 17. Lochte T. Grundriss der Entwicklung des Menschlichen Haares. Frankfurt/Main: Schöps Verlag, 1951:1-144. 18. Dawber R. Diseases of the Hair and Scalp. Oxford: Blackwell, 1997. 19. Paus R. Control of the hair cycle and hair diseases as cycling disorders. Curr Opin Dermatol 1996; 3:248-258. 20. Stenn KS, Combates N, Eilertsen KJ et al. Hair follicle growth controls. Dermatol Clinics 1996; 14:543-558. 21. Jahoda CAB, Horne KA, Oliver RF. Induction of hair growth by implantation of cultured dermal papilla cells. Nature 1984; 311:560-562. 22. Jahoda CAB, Reynolds AJ. Dermal-epidermal interactions: Adult follicle-derived cell populations and hair growth. Dermatol Clinics 1996; 14:573-583. 23. Paus R, Foitzik K, Welker P et al. Transforming growth factor-β type I and type II expression during murine hair follicle development and cycling. J Invest Dermatol 1997; 109:518-526. 24. Oliver RF. Whisker growth after removal of the dermal papilla and lengths of the follicle in the hooded rat. J Embryol Exp Morphol 1966; 15:331-347. 25. Panaretto BA. Gene expression of potential morphogens during hair follicle and tooth formation: A review. reprod fertil Dev 1993; 5:345-360. 26. Chuong C-M. The making of a feather: Homeoproteins, retinoids and adhesion molecules. BioEssays 1993; 15:513-521. 27. Chase HB. Growth of the hair. Physiol rev 1954; 34:113-126. 28. Montagna W, Ellis RA. The biology of hair growth. New York: Academic Press, 1958. 29. Straile WE. In: Lyne AG, Short BF, eds. Biology of Skin and Hair Growth. Sydney: Angus and Robertson, 1965:35-57. 30. Cotsarelis G, Sun T-T, Lavker RM. Label retaining cells reside in the bulge area of the pilosebaceous unit: Implicatios for follicular stem cells, hair cycle and skin carcinogenesis. Cell 1990; 61:1329-1337. 31. Hébert JM, Rosenquist T, Götz J et al. FGF5 as a regulator of the hair growth cycle: Evidence from targeted and spontaneous mutations. Cell 1994; 78:1017-1025. 32. Sundberg JP, ed. Handbook of mouse mutations with skin and hair abnormalities: Animal models and biomedical tools. Boca Raton Fl: CRC Press, 1994. 33. Lyons KM, Pelton RW, Hogan BM. Organogenesis and pattern formation in the mouse: RNA distribution pattern suggests a role for bone morphogenetic protein-2A (BMP-2A). Development 1990;109:833.

106

Molecular Basis of Epithelial Appendage Morphogenesis

34. Breedis C. Regeneration of hair follicles and sebaceous glands from epithelium of scars in the rabbit. Cancer Res 1954; 14:575-579. 35. Cachon-Gonzalez MB, Fenner S, Coffin JM et al. Structure and expression of the hairless gene in mice. 1994; 91:7717. 36. Billingham RE, Russell PS. Incomplete wound contracture and the phenomenon of hair neogenesis in rabbit’s skin. Nature 1956; 177:791-792. 37. Billingham RE. A reconsideration of the phenomenon of hair neogenesis with particular reference to the healing of cutaneous wounds in adult mammals. In: Montagna W, Ellis RA, eds. The Biology of Hair Growth. New York: Academic Press, 1958:451-468. 38. Fleischhauer K. Über die Entstehung der Haaranordnung und das Zustandekommen räumlicher Beziehungen zwischen Haaren und Schweissdrüsen. Z Zellforsch u mikroskop Anat 1953; 38:328-355. 39. Holbrook KA, Odland GF. Structure of the hair canal and initial hair eruption. J Invest Dermatol 1978; 71:385-390. 40. Breathnach AS, Smith J. Fine structure of the early hair germ and dermal papilla in the human fetus. J Anat 1968; 102:511-526. 41. Hashimoto K. The ultrastructure of the skin of human embryos V. The hair germ and perifollicular mesenchymal cells. Br J Derm 1970a; 83:167-176. 42. Carlsen RA. Human fetal hair follicles: The mesenchymal component. J Invest Dermatol 1974; 63:206-211. 43. Wessells NK, Roessner KD. Nonproliferation in dermal condensation of mouse vibrissae and pelage hairs. Dev Biol 1965; 12:419-433. 44. Robins EJ, Breathnach AS. Fine structure of the human foetal hair follicle at hair peg and early bulbous peg stages of development. J Anat 1969; 104:563-569. 45. Lane EB, Wilson CA, Hughes BR et al. Stem cells in hair follicles: Cytoskeletal studies. Ann NY Acad Sci 1991; 642:197-213. 46. Stasiak PC, Purkiss PE, Leigh IM et al. Keratin 19: Predicted amino acid sequence and broad tissue distribution suggests it evolved from keratinocyte keratins. J Invest Dermatol 1989; 92:707-716. 47. Montagna W, Van Scott E. The anatomy of the hair follicle. In: Montagna W, Ellis RA, eds. The Biology of Hair Growth. New York: Academic Press, 1958:39-64. 48. Robins EJ, Breathnach AS. Fine structure of bulbar end of foetal hair follicle at stage of differentiation of inner root sheath. J Anat 1970; 107:131-146. 49. Hashimoto K. The ultrastructure of the skin of human embryos VI. Formation of the intradermal hair canal. Dermatologica 1970b; 141:49-54. 50. Kerr JFR, Wyllie AH, Currie AR. Apoptosis: A basic biological phenomenon with wide ranging implications in tissue kinetics. Br J Cancer 1972; 26:239-257. 51. Smith DW, Gong BT. Scalp hair patterning: Its origin and significance relative to early brain and upper facial development. Teratology 1974; 9:17-34. 52. Serri F, Montagna W, Mescon H. Studies of the skin of the fetus and the child. J Invest Dermatol 1962; 39:199-217. 53. Barth JH. Normal hair growth in children. Pediatric Dermatology 1987; 4:173-184. 54. Kaplan ED, Holbrook KA. Dynamic expression patterns of tenascin, proteoglycans and cell adhesion molecules during human hair follicle morphogenesis. Dev Dyn 1994; 199:141-155. 55. Davidson P, Hardy MH. The development of the mouse vibrissae in vivo and in vitro. J Anat 1952; 86:342-356. 56. Handjiski B, Eichmüller S, Hofmann U et al. Alkaline phosphatase activity and localization during the murine hair cycle. Br J Dermatol 1994; 131:303-310. 57. Eichmüller S, van der Veen C, Moll I et al. J Histochem Cytochem, 1998; 46:361-370. 58. Mina M, Kollar EJ. The induction of odontogenesis in non-dental mesenchyme combined with early murine mandibular arch epithelium. Arch Oral Biol 1987; 32:123-127. 59. Hirai Y, Nose A, Kobayashi S et al. Expression and role of E- and P-cadherin adhesion molecules in embryonic histogenesis. II. skin morphogenesis.Development 1989; 105:271-277.

Principles of Hair Follicle Morphogenesis

107

60. Hirai Y, Takebe K, Takashina M et al. Epimorphin: A mesenchymal protein essential for epithelial morphogenesis. Cell 1992; 69:471-481. 61. Holbrook KA, Smith LT, Kaplan ED et al. Expression of morphogens during human follicle development in vivo and a model for studying follicle morphogenesis in vitro. J Invest Dermatol 1993; 101:39S-49S. 62. Jiang T-X, Chuong C-M. Mechanisms of skin morphogenesis. I. Analyses with antibodies to adhesion molecules tenascin N-CAM and integrin. Dev Biol 1992; 150:82-98. 63. Green MR, Couchman. Distribution of epidermal growth factor receptors in rat tissues during embryonic skin development, hair formation and the adult hair growth cycle. J Invest Dermatol 1984; 83:118-123. 64. Polakowska RR, Haake A. Apoptosis: The skin from a new perspective. Death Differentiation 1994; 1:19-31. 65. Polakowska RR, Piacentini M, Bartlett R et al. Apoptosis in human skin development: Morphogenesis, periderm, and stem cells. Dev Dyn 1994; 199:176-188. 66. Lindner G, Botchkarev VA, Botchkareva NV et al. Analysis of apoptosis during murine hair follicle regression (catagen). Am J Pathol 1997; 151:1601-1617. 67. Korsmeyer SJ. Bcl-2 initiates a new category of oncogenes: Regulators of cell death. Blood 1992; 80:879-886. 68. Stenn KS, Lawrence L, Veis D et al. Expression of Bcl-2 protoncogene in the cycling adult mouse hair follicle. J Invest Dermatol 1994; 103:107-111. 69. Chuong C-M, Oliver G, Ting SA et al. Gradient of homeoproteins in developing feather buds. Development 1990; 110:1021-1030. 70. Kanzler B, Viallet JP, Mouellic H et al. Differential expression of two different homeobox gene families during mouse tegument morphogenesis. Int J Dev Biol 1994; 38:633-640. 71. MacKenzie A, Leeming GL, Jowett AK et al. The homeobox gene Hox 7.1 has specific regional and temporal expression patterns during early murine craniofacial embryogenesis, especially tooth development in vivo and in vitro. Dev 1991; 111:269-285. 72. MacKenzie A, Ferguson M, Sharpe PT. Expression patterns of the homeobox gene, Hox-8, in the mouse embryo suggests a role in specifying tooth initiation and shape. Development 1992; 115:403-420. 73. Noveen A, Jiang T-X, Ting-Bereth SA et al. Homeobox genes Msx-1 and Msx-2 are associated with induction and growth of skin appendages. J Invest Dermatol 1995; 104:711-719. 74. Hardy MH. Glandular metaplasia of hair follicles and other responses to vitamin A excess in cultures of rodent skin. J Embryol Exp Morphol 1968; 19:157-180. 75. Hardy MH, Dhouailly D, Torma H et al. Either chicken embryo dermis or retinoid treated mouse dermis can initiate glandular morphogenesis from mammalian epidermal tissue. J Exp Zool 1990; 256:279-289. 76. Viallet JP, Dhouailly D. Retinoic acid and mouse skin morphogenesis. I. Expression pattern of retinoic acid receptor genes during hair vibrissa follicle, plantar, and nasal gland development. J Invest Dermatol 1994; 103:116-121. 77. Ruberte E, Dolle P, Krust et al. Specific spatial and temporal distribution of retinoic acid receptor gamma transcripts during mouse embryogenesis. Dev Biol 1990; 108:213-222. 78. Muller WA. Developmental Biology. NY: Springer, 1997. 79. Hogan BLM. Bone morphogenic proteins: Multifunctional regulators of vertebrate development. Genes and Development 1996; 10:1580-1594. 80. Roberts A, Sporn M. The transforming growth factor-βs. In: Sporn M, Roberts A, eds. Peptide Growth Factors and Their Receptors (Handbook of Experimental Pharmacology, Vol 95). New York: Springer-Verlag, 1990:419. 81. Pelton RW, Nomura S, Moses HL, Hogan BM. Expression of transforming growth factors β 2 during murine embryogenesis. Development 1989; 106:759-767. 82. Pelton RW, Saxena B, Jones M et al. Immunohistochemical localization of TGF β1, TGF β2 and TGF β3 in the mouse embryo: Expression patterns suggest multiple roles during embryonic development. J Cell Biol 1991; 115:1091-1105. 83. Shull MM, Ormsby I, Kier AB et al. Targeted disruption of the mouse transforming growth factor-B1 gene results in multifocal inflammatory disease. Nature 1992; 359:693-699.

108

Molecular Basis of Epithelial Appendage Morphogenesis

84. Kulkami AB, Ward JM, Yaswen L et al. Transforming growth factor beta1 null mice. An animal model for inflammatory disorders. Am J Pathol 1993; 146:264. 85. Philpott MP, Green MR, Kealey T. Human hair growth in vitro. J Cell Sci 1990; 93:409418. 86. Jones CM, Lyons KM, Hogan BM. Involvement of bone morphogenic protein-4 (BMP-4) and Vgr-1 in morphogenesis and neurogenesis in the mouse. Development 1991; 111:531-542. 87. Vainio S, Karavanova I, Jowett A et al. Identification of BMP-4 as a signal mediating secondary induction between epithelial and mesenchymal tissues during early tooth development. Cell 1993; 75:45-58. 88. Blessing M, Nanney LB, King LE et al. Transgenic mice as a model to study the role of TGF-β-related molecules in hair follicles. Genes and Dev 1993; 7:204-215. 89. Takahashi H, Ikeda T. Transcripts for two members of the transforming growth factor-β superfamily BMP-3 and BMP-7 are expressed in developing rat embryos. Dev Dyn 1996; 207:439-449. 90. Goodrich LV, Johnson RL, Milenkovic L et al. Conservation of the hedgehog/patched signaling pathway from flies to mice: Induction of a mouse patched gene by hedgehog. Genes and Dev 1996; 10:301-312. 91. Ting-Bereth SA, Chuong C-M. Sonic hedgehog in feather morphogenesis: Induction of mesenchymal condensation and association with cell death. Dev Dyn 1996; 207:157-170. 92. Nanney LB, Stoscheck CM, King LE et al. Immunolocalization of epidermal growth factor receptors in normal developing human skin. J Invest Dermatol 1990; 94:742-748. 93. Philpott MP, Kealey T. The effects of EGF on the morphology and patterns of DNA synthesis in isolated human hair follicles. J Invest Dermatol 1994; 102:186-191. 94. Moore GPM, Panaretto BA, Robertson DJ. Effects of epidermal growth factor on hair growth in the mouse. J Endocrinol 1981; 88:293-299. 95. Moore GPM, Panaretto BA, Robertson DJ. Epidermal growth factor delays the development of the epidermis and hair follicles of mice during growth of the first coat. Anat Rec 1983; 205:47-55. 96. Luetteke NC et al. TGF-α deficiency results in hair follicle and eye abnormalities in targeted and waved-1 mice. Cell 1993; 73:262-278. 97. Luetteke NC et al The mouse waved-2 phenotype results from a point mutation in the EGF receptor tyrosine kinase. Gene Dev 1994; 8:399-423. 98. Peus D, Pittelkow MR. Growth factors in hair organ development and the hair growth cycle. Dermatologic Clinics 1996; 14:559-572. 99. Threadgill DW, Dlugosz AA, Hansen LA et al. Targeted disruption of mouse EGF receptor: Effects of genetic background on mutant phenotype. Science 1995; 269:230-237. 100. Gruneberg H. The glandular aspects of the tabby syndrome in the mouse. J Embryol Exp Morphol 1971; 25:1-19. 101. Mayer TC, Green MC. Epidermis is the site of action of Tabby (Ta) in the mouse. Genetics 1978; 90:125-131. 102. Claxton JH. The initiation and development of the hair follicle population in tabby mice. Genet Res 1967; 10:161-171. 103. Kapalanga J, Blecher SR. Effect of the X-linked gene Tabby (Ta) on eyelid opening and incisor eruption in neonatal mice is opposite to that of epidermal growth factor. Development 1990; 108:349-355. 104. Blecher SR, Kapalanga J, Lalonde D. Induction of sweat glands by epidermal growth factor in murine X-linked anhydrotic ectodermal dysplasia. Nature 1990; 345:542-545. 105. Widelitz RB, Jiang T-X, Noveen A et al. FGF induces new feather buds from developing avian skin. J Invest Dermatol 1996; 107:797-803. 106. Werner S, Smola H, Liao X et al. The function of KGF in the morphogenesis of epithelium and repithelialization of wounds. Science 1994; 266:819-822. 107. Guo L, Degenstein L, Fuchs E. Keratinocyte growth factor is required for hair development but not for wound healing. Genes Dev 1996; 10:165-175.

Principles of Hair Follicle Morphogenesis

109

108. Danilenko DM, Ring BD, Yanagihara D et al. Keratinocyte growth factor is an important endogenous mediator of hair follicle growth development and differentiation. Am J Pathol 1995; 147:145-154. 109. Peters KG, Werner S, Chen G et al. Two FGF receptor genes are differentially expressed in epithelial and mesenchymal tissues during limb formation and organogenesis in the mouse. Development 1992; 114:233-243. 110. Rosenquist TA, Martin GR. Fibroblast growth factor signaling in the hair growth cycle: Expression of the fibroblast growth factor receptor and ligand genes in the murine hair follicle. Dev Dyn 1996; 205:379-386. 111. Du Cros DL. Fibroblast growth factor and epidermal growth factor in hair development. J Invest Dermatol 1993; 101:101S-106S. 112. DuCros DL, Isaacs K, Moore GPM. Distribution of acidic and basic fibroblast growth factors in ovine skin during hair follicle morphogenesis. J Cell Sci 1993; 105:667-674. 113. Finch PW, Cunha GR, Rubin JS et al. Pattern of keratinocyte growth factor and keratinocyte growth factor receptor expression during mouse fetal development suggests a role in mediating morphogenetic mesenchymal-epithelial interactions. Dev Dyn 1995; 203:223-240. 114. Niswander L, Martin G. FGF-4 expression during gastrulation, myogenesis, limb and tooth development in the mouse. Development 1992; 114:755-768. 115. Mercola M, Stiles CD. Growth factor superfamilies and mammalian embryogenesis. Development 1988; 102:451-460. 116. Liu J-P, Baker J, Perkins AS et al. Mice carrying null mutations of the genes encoding insulin-like growth factor I (IGF-I) and type 1 IGF receptor (Igf1r). Cell 1993; 75:59-72. 117. Bol DK, Kiguchi K, Gimenez-Conti I et al. Overexpression of insulin-like growth factor-1 induces hyperplasia, dermal abnormalities, and spontaneous tumour formation in transgenic mice. Oncogene 1997; 14:1725-1734. 118. Zhou P, Byrne C, Jacobs J et al. Lymphoid enhancer factor 1 directs hair follicle patterning and epithelial cell fate. Genes Dev 1995; 9:570-583. 119. Kratochwil K, Dull M, Farinas I et al. Lef1 expression is activated by BMP-4 and regulates inductive tissue interactions in tooth and hair development. Genes Dev 1996; 10:1382-1394. 120. Brissette JL, Li J, Kamimura J et al. The product of the mouse nude locus, Whn, regulates the balance between epithelial cell growth and differentiation. Genes Dev 1996; 10:2212-2221. 121. Kopf-Maier P, Mboneko VF, Merker HJ. Nude mice are not hairless. Acta Anat 1990; 139:178-190. 122. Panteleyev AA, van der Veen C, rosenbach T et al. Towards defining the pathogenesis of the hairless phenotype. J Invest Dermatol 1998; 110:902-907. 123. Fuchtbauer EM. Expression of M-Twist during postimplantation development of the mouse. Dev Dyn 204:316-322, 1995. 124. Jen Y, Manova K, Benezera R. Expression patterns of Id1, Id2 and Id3 are highly related but distinct from that of Id4 during mouse embryogenesis. Dev Dyn 207:235-252, 1996. 125. Mitsiadis TA, Salmivirta M, Muramatsu T et al. Expression of the heparin binding cytokines, midkine (MK) and HB-GAM (pleiotrophin) is associated with epithelial-mesenchymal interactions during fetal development and organogenesis. Dev 1995; 121:37-51. 126. Mitsiadis TA, Henrique D, Thesleff I et al. Mouse serrate-1 (jagged-1); expression in the developing tooth is regulated by epithelial-mesenchymal interactions and fibroblast growth factor-4. Development 1997; 124:1473-1483. 127. Weinmaster G, Roberts VJ, Lemke G. A homologue of Drosophila Notch expressed during mammalian development. Development 1991; 113:199-205. 128. Kopan R, Weintraub H. Mouse Notch: expression in hair follicles correlates with cell fate determination. J Cell Biol 1993; 121:631-641. 129. Reference deleted in proof. 130. Piepkorn M, Underwood, Henneman C et al. Expression of amphiregulin is regulated in cultured human keratinocytes and in developing fetal skin. J Invest Dermatol 1995; 105:802. 131. Lane TF, Deng C, Elson A et al. Expression of Brca1 is associated with terminal differentiation of ectodermally and mesodermally derived tissues in mice. Gene Dev 1995; 9:2712.

110

Molecular Basis of Epithelial Appendage Morphogenesis

132. Seiberg M, Marthinuss J. Clusterin expression within skin correlates with hair growth. Dev Dyn 1995; 202:294-301. 133. Risek B, Klier G, Gilula NB. Multiple gap junction genes are utilised during rat skin and hair development. Dev 1992; 116:639-651. 134. Kaya G, Rodriguez I, Jorcano JL et al. Selective suppression of CD44 in keratinocytes of mice bearing an antisense CD44 transgene driven by a tissue-specific promoter disrupts hyaluronate metabolism in the skin and impairs keratinocyte proliferation. Genes Dev 1997; 11:996-1007. 135. Bitgood MJ, McMahon AP. Hedgehog and BMP genes are coexpressed at many diverse sites of cell-cell interaction in the mouse embryo. Dev Biol 1995; 172:126-138. 136. Iseki S, Araga A, Ohuchi H et al. Sonic hedgehog is expressed in epithelial cells during development of whisker, hair and tooth. Biochem Biophys Res Com 1996; 218:688- 693. 137. Bondy C, Werner H, Roberts CT et al. Cellular patterns of insulin-like growth factor-I (IGF-I) and type-I IGF receptor gene expression in early organogenesis; comparison with IGF-II gene expression. Mol Endocrinol 1990; 4:1386-1398. 138. Sonnenberg E, Meyer D, Weidner KM et al. Scatter factor/hepatocyte growth factor and its receptor the c-met tyrosine kinase, can mediate a signal exchange between mesenchyme and epithelia during mouse development. J Cell Biol 1993; 123:223-235. 139. Davies AM, Bandtlow C, Heumann R et al. Timing and site of nerve growth factor synthesis in developing skin in relation to innervation and expression of the receptor. Nature 1987; 326:353. 140. Holbrook KA, Fisher C, Dale BA et al. Morphogenesis of the hair follicle during the ontogeny of human skin. In The biology of wool and hair (Rogers GE, reis PJ, Ward KA, Marshall RC eds). London: Chapman Hall 1989:15-35. 141. Schmid P, Lorenz A, Hameister H et al. Expression of p53 during mouse embryogenesis. development 1991; 113:857. 142. Botchkarev NV, Albers KM et al. Neurotrophin -3 involvement in the regulation of hair follicle morphogenesis. J Invest Dermatol 1998; in press. 143. Vassalli A, Matzuk MM, Gardner HAR et al. Activin/inhibin bB subunit gene disruption leads to defects in eyelid development and female reproduction. Genes Dev 1994; 8:414. 144. Guo L, Yu Q-C, Fuchs E. Targeting expression of keratinocyte growth factor to keratinocytes elicits striking changes in epithelial differentiation in transgenic mice. EMBO J 1993; 12:973. 145. Matzuk MM, Lu N, Vogel H et al. Multiple defects and perinatal death in mice deficient in follistatin. Nature 1995; 374:360. 146. Turksen K, Kupper T, Degenstein L et al. Interleukin 6: Insights to its function in skin by overexpression in transgenic mice. Proc Natl Acad Sci USA 1992; 89:5068. 147. Wysolmerski JJ, Broadus AE, Zhou J et al. Overexpression of parathyroid hormone- related protein in the skin of transgenic mice interferes with hair follicle development. Proc Natl Acad Sci USA. 1994; 91:1133. 148. Vassar R, Fuchs E. Transgenic mice provide new insights into the role of TGF α during epidermal development and differentiation. Gen Dev 1991; 5:714. 149. Selheyer K, Bickenbach JR, Rothnagel JA et al. Inhibition of skin development by overexpression of transforming growth factor beta-1 in the epidermis of transgenic mice. Proc natl Acad Sci USA 1993; 90:5237. 150. Carroll JM, Romero R, Watt FM. Suprabasal integrin expression in the epidermis of transgenic mice results in developmental defects and a phenotype resembling psoriasis. Cell 1995; 83:957. 151. Dealey CN, Kosher RA. IGF-I and insulin in the acquisition of limb-forming ability by the embryonic lateral plate. Dev Biol 1996; 177:291-299. 152. Dealey CN, Clarke K, Scranton V. Ability of FGFs to promote the outgrowth and proliferation of limb mesoderm is dependent on IGF-I activity. Dev Dyn 1996; 206:463-469.

CHAPTER 7

Growth of the Hair Follicle: A Cycling and Regenerating Biological System Kurt Stenn, Satish Parimoo and Stephen M. Prouty

Introduction General

H

air growth involves the unique phenomenon of cyclic regeneration: This structure forms, regresses, and regrows over a recurring period. After a brief description of cycling and regeneration in animal biology, and the introduction of an antler cycle, in this essay we will review the hair cycle with emphasis on our current understanding of its controls. Where relevant we will allude to other cycling and regenerating systems in search of common themes. It is our message that the hair follicle is unique among mammalian tissues and among other cycling systems in that it forms, regresses and regrows throughout the lifetime of the individual. Its controls, as yet poorly defined, implicate cyclic epithelial-mesenchymal interactions, expression of growth factors, cytokines, cellular infiltrates, and immune changes. The site and basis for hair growth cycling control remain unknown. As we consider the antler and hair cycles, we are restricting our focus to a unique type of cycle. We are not focusing on the cell cycle per se, although the cell cycle is undoubtedly important to any growing biological structure. We are not focusing on the short daily cycle nor are we focusing only on a hormonal cycle, as not all follicles are sex hormone dependent. We are instead looking at a multicellular system that cyclically regenerates over a period of months to years.

Cycling Reflecting the celestial and terrestrial environment, biological rhythms can be divided into three broad classes based on their periodicity: circadian, of about 24 h; ultradian, of less than 24 h; and infradian, of greater than 24 h. These rhythms are ubiquitous and examples can be found at all levels of biological organization: 1. the cell division cycle of the prokaryotes, such as Synechococcus1 and eukaryotic cells in culture;2 2. the cell division cycles of tissue cells such as skin,3 gut,4 cornea,5 liver and bone marrow;2 3. the cycling of organs such as the hair follicle,6,7 endometrium, and deer antler;8 4. organ cycling such as the female reproductive system, endocrine secretion, and mating behavior; and 5. population growth and behavioral cycles such as the circannual mass migration of some oceanic organisms. Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

112

Molecular Basis of Epithelial Appendage Morphogenesis

The implications of rhythm have extended well beyond the phenomena of behavior prediction and reproduction, for it is recognized that tissue rhythms affect disease expression and therapeutic response.9 Here, then, is a potential advantage in the timing of metabolic, developmental and behavioral phenomena to the environment. Of these various types of cycles, one of the best studied is the 24 h circadian rhythm (reviewed in refs. 10-12). Several features are universal for circadian rhythms. They generally have three main components: an oscillator or clock that generates the rhythm, an input pathway to the oscillator that allows entrainment of the clock to the environment, and output pathways that couple the oscillation of the clock to the effector systems. The nature of the oscillator is a cell-autonomous clock based on a negative feedback loop between rhythmically-expressed mRNAs and their auto-inhibitory protein products; the best studied are the frequency gene of Neurospora, the period and timeless genes of Drosophila,12 and the Clock gene of the mouse.13 Cellular sites of the oscillators include the Neurospora cell, the Drosophila central nervous system, the retina of mammals, amphibia and mollusks, the pineal gland of bird and mammals, and the mammalian suprachiasmatic nucleus. Perturbation of signaling pathways indicate that transduction of the light signal into the oscillator cells may involve Ca2+ fluxes followed by kinase and phosphatase cascades. Other defining features of the circadian rhythm include: 1. light as the synchronizing environmental input; 2. persistence of the rhythm in the absence of environmental cues; 3. resetting of rhythm by brief environmental stimuli; and 4. temperature-independence. That the cycle is often entrained to light directly relates biological photodetection systems to the photoperiod systems. This notion has been corroborated recently by the recognition of sequence similarity in some genes of both systems.14

Regeneration Cyclic growth of any biologic structure, at any level—cellular, tissue, organ, organismal— implies its formation, regression and reformation over time. In this discussion we are distinguishing regeneration from repair. Repair would occur in a tissue unable to reform itself, and thus would replace lost tissue with scar. On a macroscale the fact that each living unit generates another of its own kind can be viewed as a sort of cyclic regeneration inherent in all life forms. More primitive animals have the additional capacity to reform body regions, large organ systems, or appendages either cyclically, such as the integument in the insect or reptile, or incidentally, after tissue loss, as limb regeneration in the amphibian.15 For higher organisms regenerative properties are, in general, limited to reproduction and not to isolated parts of the whole. It is recognized, however, that even the most developed animals retain some regenerative ability in many tissues in the fetal state and in a few restricted tissues in the adult. Regeneration may result after an injurious insult or a controlled process of autotomy.16 Complete recovery would entail the total replacement or regeneration of a missing segment, organ, tissue, or cell. In general, tissues vary in their ability to regenerate after injury or loss; for example, while blood cells and epithelial linings of the gut and skin have great regenerative ability, neural tissues do not. More relevant to our discussion here are other tissues which regenerate not as the result of tissue damage but as a result of the process of autotomy, the controlled loss of a biological structure. For the tissues under consideration the mature organ undergoes controlled atrophy, death, and shedding followed by reformation of the structure in preparation for the next autotomy signal. The antler growth cycle, the endometrial cycle and the hair growth cycle each manifest true regeneration of the original structure in response to a process of cyclic autotomy. Below we consider the antler and the

Growth of the Hair Follicle

113

hair growth cycle. The estrus cycle has been reviewed elsewhere.17 In these examples, there is a rhythm to both processes, atrophy (autotomy) and regeneration. What do regenerative systems share? Considerable investigative attention has been given to the mechanism of body segment or limb regeneration in lower animal forms, with the hope that this understanding will lead to some means of promoting regeneration in humans. In appendage regeneration, the tissue proximal to the injury undergoes a process of dedifferentiation whereby the tissue loses its differentiated character and becomes more embryonic. The dedifferentiated cells at the site of injury aggregate to form a rounded mass, the blastema, which is endowed to reform the lost structure. Certain cellular features are required by many regenerating systems145: 1. Cells which are capable of forming a blastema must be left behind after the injury. 2. A wound environment must be present. As a blastema never forms in the absence of epidermal wounding, it is assumed that a wounded epidermal cap induces the formation of the blastema beneath it. In the laboratory mouse and in children below 5 years of age,18 regeneration of an amputated finger tip will occur only if the wound is not covered by an intact epidermis. 3. A supply of vessels and nerves must be present in the regenerating area.

Antler Formation8 Of the several dozen species of deer in the world all but two possess antlers. Because of their majesty, their limitation—in general—to one sex, and their occurrence during the mating season, antlers are thought to serve the purpose of sexual communication. In contrast to horns, which arise in the skin and do not cycle, antlers grow from the underlying bony structures and do cycle. When the antler sheds, it leaves an ulcer over its attachment site. In this area of “wound” the new antler arises. Antler formation initiates within periosteal tissue at a thickening of the skull referred to as the pedicle. The growth from the pedicle consists of bony mesenchyme surrounded by skin, which itself, with its hair appendages, forms de novo each cycle. Next to their regenerative ability the most unique attribute of antlers is their unusually rapid rate of elongation; it is contended that no other tissue in the animal kingdom grows as fast as antlers. When growth is complete, the bony structure of the antler hardens and the surrounding skin sheds. At the end of the mating season antler dyshesion, an active process which is exquisitely controlled and probably involves the action of osteoclasts, is initiated. Upon shedding, the antlers expose, once again, an open wound. After old antlers have been cast, skin migrates over the stump of the bony pedicle carrying both epidermal and mesenchymal tissues. This resurfacing pedicle skin shows complete regeneration: It does not form a scar. Histologically, the epidermis growing over the pedicle shows marked hyperplasia reminiscent of that overlying an embryonic limb bud or the blastema of a regenerating limb. In order to effect normal antler regeneration, the dermis of this regenerating skin must contain normal vascular and nerve supplies as well as the blastema. A given antler growth cycle progresses over the course of a year, but the cycles continue throughout the lifetime of the buck. Antler growth control is related to the mating cycle of deer. This cycle is entrained to the annual light cycle as an environmental cue to the time of year. The light cycle affects the reproductive cycle and the fur molting cycle, as well as the antler growth cycle of these animals. The stimulus for antler replacement is entirely dependent on the fluctuations of day length. Antler growth is stimulated when the ratio of light to dark periods over 24 h does not equal one and is suppressed when the ratio is one (the lengths of the day and the night are equal). The photoperiod message is processed by the eye and transmitted to the suprachiasmatic nuclei, which, in turn, neuronally signal the pineal

114

Molecular Basis of Epithelial Appendage Morphogenesis

gland through sympathetic innervation.19 Substances synthesized by the pineal gland coordinate the reproductive activity associated with the photoperiod and thus the rhythm of the antler cycle. If the pineal gland is ablated, antler growth and the casting schedules go awry; the bucks cast their antlers in spring instead of autumn and their winter pelage, which is usually shed in spring, is retained into the summer. The message relating the environmental events is carried by circulating androgenic hormones to the end organs. Antlers are thus secondary sex characteristics and as such are primarily affected by the annual fluctuations in the secretion of sex hormones. The hormone acts on the periosteum of the pedicles and stimulates the deposition of bone. With the production of high levels of testosterone, bone formation ceases, ossification strengthens the bone formed, and the skin enwrapping the antler bone sheds. After the mating season, testosterone levels drop and the antler sheds. In contrast to high levels of testosterone, when estrogens are given to castrated bucks antler shedding is delayed. The growth of antlers is independent of the shedding of the old growth before the new appears; that is, the loss of the old antler and growth of the new are not causally related. It appears that declining concentrations of male hormones are responsible for the shedding and the initiation of new antler growth. However, wound healing would appear to be important to new antler formation, since antler growth begins with the healing of the raw surface of the stump and, if the stump is closed over with full-thickness skin grafts, antler regeneration is prevented altogether.8 Antlers will develop only where a wound is allowed to heal on the pedicle. These reparative/regenerative features can also be seen in other morphogenetic systems such as in tooth formation, regenerating amphibian limb, and feather growth in the bird (some of these topics are considered elsewhere in this volume). Antlers are relevant to the following discussion because: 1. they undergo a long growth cycle over the period of a year; 2. growth and shedding of antlers appear to be independent events; 3. wound environment is important to antler regrowth; 4. antler growth and shedding is hormonally determined. They stop growing to maturity under conditions of high testosterone levels; antler shedding is delayed by high concentrations of estrogen; 5. complete antler regeneration is observed at the end of each cycle over the lifetime of the animal.

Cycle of the Hair Follicle Like Antler Development, Hair Growth Is Cyclic The hair follicle undergoes a cycle that involves periods of growth (anagen), regression(catagen), rest (telogen), and shedding20 (Fig. 7.1). Because of his dependence on animal fur,15,21 man has long recognized that wool has periods of growth and shedding. In most animals hair grows synchronously in waves spreading over the animal skin surface in recognizable patterns; in man, however, each follicle grows asynchronously, apparently independent of its neighbor. Where studied, the growth cycle of fur is regulated by the seasonal changes in the length of the day, as in the case of the antler entrained to the photoperiod.22 In mink, for example, increasing day length in the spring induces growth of the summer fur and molting of the winter fur; in contrast, decreasing day length in the fall initiates growth of the winter fur and molting of the summer fur. Ablation studies have led to the notion that the annual cycle of fur growth is regulated in mink and sheep23 by a neuroendocrine mechanism involving the pineal gland and the pineal hormone melatonin. That winter coat growth can be induced by melatonin suggests that a short photoperiod is the stimulus for melatonin secretion by the pineal gland. Melatonin binding sites have been

Growth of the Hair Follicle

115

Fig. 7.1. Hair growth cycle: Stages of follicle. The hair follicle is conceived of as a cycle with an entrance limb giving the cycle the shape of a figure “6”. After morphogenesis, which occurs only once, the follicle cycles throughout life, in defined stages. The dotted-line-cycles refer to those follicles which are androgen sensitive. While the stages are drawn here with defined periods, in fact the stages of follicle growth vary from follicle to follicle.

found on the epithelial bulb of the follicle.24 It has been suggested that melatonin acts on the follicle in part through the action of prolactin; moreover, prolactin secretion is under the control of melatonin.25,26 A decrease in circulating prolactin coincides with the development of a winter coat, and injections of prolactin block the development of that coat in mink.27 Interestingly, it has been found that there is a circannual rhythm imposed on the circadian rhythm of prolactin secretion in the red deer, in that blood prolactin concentrations reach a maximum during the nonbreeding season.28 That wool production is affected by the photoperiod, and that this effect can be separated from nutrition and temperature,15 is of potential significance to the wool production industry. In the laboratory, it has been demonstrated that hair follicles may cycle independently of environmental signals. The cycles for these follicles appear to be imprinted and internalized, and they will continue to act for some time in the absence of external stimuli, for example, after follicle transplantation. However, if the external stimuli begin to change, the internal clock of the follicle will adjust itself to come into line with the changed conditions. In studies with rats, Ebling & Johnson29 found that the hair follicles of transplanted skin

116

Molecular Basis of Epithelial Appendage Morphogenesis

continue to maintain periodicity characteristic of their sites of origin. Using parabiotic animals, however, they were able to show that, with time, hair growth tended to become synchronized. This observation was interpreted to indicate that inherent rhythms are slowly brought into phase by a systemic factor,30 but that systemic factors do not control the cycle intimately; so, according to these workers the cyclic activity of the follicle depends upon an inherent rhythm which is able to respond slowly to an undefined systemic stimulus.31 Additional support for the conclusion that there is some intrinsic factor orchestrating the cycle was provided by Durward and Rudall,32 who found that upon rotating skin growth waves progressed as they would have in the original nonrotated position. For most animals, then, there are probably different levels of control of the hair cycle: (1) intrinsic, (2) systemic, and (3) environmental.33 Relevant to animal husbandry, the first two factors have been found to be genetic. Whether that signal is transmitted from cell to cell by means of an intercellular wave or by means of a soluble factor(s) is not known. Although many molecules have been examined for an expression pattern over the hair cycle,6,34,35 none described to date fit the pattern of that shown for the Neurospora or Drosophila clock. However, in these simpler systems, it has been shown that pulses of transcription and translation inhibitors can disrupt the rhythm when given at a specific time.36 A similar experiment could be carried out during the mouse hair cycle. Mutational analysis has provided a vital tool for identification of the clock components in fungi, flies, and mouse. Two mammals have mutations that disrupt the circadian rhythm: murine clock and hamster tau .13,19,37 The rhythmic control of human hair is less well understood. The hair growth begins in utero within the third month of gestation.38 During fetal life and the 10 weeks following birth, hair grows uniformly over the scalp.39 The wave progresses along a frontal to occipital direction. From the 18th week after birth onward, follicles progressively adopt the mosaic, and asynchronous, pattern characteristic of adults.40 Thereafter, the first scalp cycle is short, lasting less than 9 months, while the cycle of the mature scalp follicle lasts from 2 to 6 years. Important to the asynchrony of hair growth in humans is the heterogeneity of hair over the body (actually a feature of all mammals). Hairs differ greatly from site to site41 and, with the exception of bilateral symmetry, there are probably no two identical hairs on the body. Hairs differ in length, color, curl, texture and androgen sensitivity. Hairs in the central axilla, for example, have a greater rate of growth and density than in the peripheral axilla.42 Hair density does not differ between males and females, but the type of hair and growth rate does differ.43 Basic to the morphological heterogeneity of hair is the asynchrony of the hair cycle. Many animals, then, have a seasonal pattern of hair coat replacement wherein the follicles are dormant in the winter (telogen) and active during the summer (anagen) after a spring molt. In these animals the annual cycle is controlled by day length. The shortening day in autumn triggers follicle growth cessation (catagen) leading to the winter resting period, while the lengthening day in spring stimulates regrowth of the follicle with the formation of the new coat. The new coat forms before the old is lost, which assures that the animal is never exposed or naked to the environment. It is interesting that circannual rhythms have been noted in humans as well.44,45 The proportion of follicles (beard and thigh) in the growth phase, anagen, reach a single peak of over 90% in March; thereafter, the percent of growing follicles decreases to a nadir in September. The number of shed hairs peaks in late summer (August/September). How the environment influences human hair growth and what in the environment induces these changes are not yet clear.

Growth of the Hair Follicle

117

Like Antler Development Hair Growth Is Regenerative A past and current criticism of many investigative studies of skin physiology is that the influence of the hair cycle on the phenomenon studied is usually disregarded.46 It has been demonstrated many times that the hair cycle imposes cyclic and profound changes on the morphology and function of the total skin organ, as well as the pilosebaceous apparatus.46-49 Throughout the cycle there are changes in the tissues, cells, and molecules that make up the skin; moreover, the skin reacts to environmental stimuli differentially depending on the cycle phase (e.g., ref. 50). Imposed on the hair cycle is a circadian rhythm of the epidermal cells, and, although not yet demonstrated, probably a similar rhythm of the mesenchymal cells.2 The hair cycle and the molecules associated with it have recently been reviewed.6,7 As emphasized in those reviews, although we currently recognize molecular associations with certain phases of the cycle, with few exceptions we do not know what actual role these molecules play in initiating, supporting, or terminating the cycle. The morphology of the hair cycle was defined first by Dry20 who recognized growth, regression, resting and shedding phases of that cycle. The individual phases of the cycle have been further subdivided into 6 to 8 stages of anagen46,51 and 8 stages of catagen.52

Anagen The formation of a new inferior portion of the hair follicle, initiated by the cells of the resting hair germ in each cycle, is the only case of spontaneous organogenesis that occurs repeatedly throughout the lifetime of the organism from infancy to senility. Anagen (Fig. 7.2) is the phase of hair follicle growth. Relevant to that growth are several important features: 1. Follicle growth depends on critical mesenchymal to epithelial and epithelial to mesenchymal interactions. The follicular papilla, the mesenchymal nubbin at the base of the anagen follicle, consists of cells that have the ability to induce hair follicles heterotopically. 2. Hair follicle product, a shaft, is the result of highly specialized epithelial differentiation. The shaft consists of tightly bound keratinized epithelial cells. 3. Follicle differentiation involves the formation of concentric, distinct, cylindrical epithelial layers which make up two compartments: the centrally lying shaft embedded in the internal root sheath, and an outermost outer root sheath, which separates the whole structure from the dermis. 4. An associated pigmentation function is closely entrained to the follicle growth cycle. The pigmentation cycle begins after shaft formation starts and ends before shaft formation ceases, so that the shaft has an unpigmented tip and root. Although we will not develop this aspect of the hair follicle cycle here, we would like to emphasize that the pigmentation cycle is coordinated with the hair growth cycle and that the pigmentation cycle and the constituent melanocytes may have a significant influence on follicle growth. 5. Androgen-sensitive follicles, found regionally, have the potential to switch, upon androgen exposure, from a small, vellus follicle to a large, pigmented, terminal follicle. Although we know roughly how long a follicle lasts in anagen in a given region and given animal,41 we do not know what controls the onset of spontaneous anagen nor from whence the stimulus for initiating that anagen begins. Anagen can be induced artificially in the laboratory by various types of traumatic insult (plucking, wounding, vigorous shaving, topical irritants, etc.).53-56 It has been found experimentally that a certain threshold of injury is necessary to induce hair growth; for plucking that stimulus is roughly 1000 hair follicles.57 The trauma/wound healing environment that initiates anagen is reminiscent of

118

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 7.2. Hair growth cycle: Stages of shaft. This figure illustrates the transformation of the follicle and its shaft over the stages of the follicle. The figure is meant to illustrate that the course of a given shaft is different from the course of a given follicle (Adapted from Montagna and Parakkel144 by Jane Stephens).

the wound environment supporting antler growth initiation (see above), limb regeneration and trauma-induced regeneration of muscle and bone. That trauma induces anagen led early workers in the field to suggest that the hair cycle may be controlled by release from inhibition.46,58 It was Bullough who first proposed the notion that wounding reduces the concentration of an inhibitor.59 In a provocative study, Paus et al60 presented evidence that telogen epidermis contains a factor that slows or blocks anagen induction. The opposite conclusion was drawn by Johnson and Ebling,61 who contended that an inductive stimulus is released by plucking. Hormones play a role in anagen induction. For example, thyroid hormones are seen to accelerate the onset of anagen. The hormones of pregnancy and birth control pills result in an increase in the proportion of follicles in anagen, apparently delaying the onset of catagen.62 The release of this anagen stimulus leads to the commonly observed postpartum hair loss (telogen effluvium).63,64 Finally, glucocorticoids may suppress anagen induction.65 Molecules stimulating various metabolic or control pathways have also been tested for their anagen inductive ability. Such active molecules include cyclosporine A,66 FK506,67 topical minoxidil,68 substance P,69 HGF70 and KGF (FGF-7).71 From where and upon which cell the spontaneous anagen signal is sent and received is unknown. A widely held hypothesis—“the bulge activation hypothesis”72—states that the signal for anagen initiation arises in the follicular papilla73 and acts upon a nest of cells with

Growth of the Hair Follicle

119

stem cell-like properties in the follicular bulge.74-77 Not incompatible with these data, however, is the notion that the initiating signal arises within the follicular epithelium and acts back upon the resting papilla. Once the signal for follicle growth initiation is given to the resting follicle, the lower follicle starts to grow by invading the dermis and traveling along a pathway, the fibrous stele, traversed by every growth cycle of that follicle from the first. That event would appear to require synthesis of extracellular matrix and the controlled release of proteases.78,79 With the initiation and development of anagen, a series of growth factors and structural genes are alternatively transcribed and silenced.6,34 During anagen, the upper follicular canal and the sebaceous gland play an important role in processing the mature shaft.80 Reminiscent of a yo-yo, the only cells that truly cycle are the papilla cells that move up and down with the follicle from the resting, to the growing, and then back up to the resting follicle. The papilla presents as a cluster of tightly packed cells in telogen, closely adjacent to the resting follicle germ cells. The papilla accumulates significant extracellular matrix during anagen, and then with the onset of catagen again loses matrix and condenses.81,82 Since the cells of the papilla are not believed to divide during the cycle,83,84 the papilla appears to be the one fixed element of the cycling follicular structure. Continued and constant contact of the follicular epithelium with the papilla is necessary throughout the cycle.85 Once anagen is initiated the follicle differentiates and transforms through recognized stages of maturation.46,51 The bulb or matrix differentiates to give rise to 8 cell lineages that produce the differentiated products: the sheaths, the slippage plane, the cuticles, and the shaft.86,87 The mechanism of the movement of the shaft upward is variously ascribed to growth pressure from the rapidly dividing matrix or from the more rapid upward movement of the outer root sheath. Another idea not yet fully tested is that a force resulting from condensing cells of the proximal keratinizing shaft draws the structure outward by a summation of all the apoptotic cells in this region.88 This force is referred to below as an “apoptotic force”(see below).

Catagen At the completion of anagen, the length of which varies with the location of the follicle, a signal is given, anagen ceases and the regressive phase of catagen begins. The steps of catagen formation that describe the follicular regression have been traced morphologically and subdivided into phases by Straile et al.52 The earliest sign is the cessation of growth within the matrix of the follicle, followed by loosening and atrophy of the papilla, and then regression of the lower follicular epithelium. At this time the immunoreactivity of the follicle returns.89,90 The deep follicular epithelium, previously MHC class I negative, now becomes positive and the follicle base is surrounded by macrophages, cells not found about the growing follicle. At the end of the catagen phase follicular remnants are cleaned up by these cells.91 The important process in accomplishing catagen is the regression of the inferior, cycling portion of the follicle. As in other sculpting biological systems, this process of regression involves apoptosis.92 While apoptotic changes are not limited to the catagen follicle, they are most prominent therein. In catagen, a number of apoptotic cells show a gradient: apoptotic cells are found at greatest concentration within the deepest (most proximal) follicle segment.93 One of the gnawing issues in hair biology is the mechanism of follicle regression upward in catagen. The process is complex and unique. The literature contains two major theories. The first contends that the club hair is pushed upward by the elongating cell column beneath it. The second contends that the follicle is forced upward (tooth paste tubelike) from pressures of the vitreous membrane94 or from shortening and squeezing upward

120

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 7.3. Theory of apoptotic force. Illustrated in this figure is the atrophy and regression of the interdigital web, starting at the upper left. A force is envisioned to develop in the web as epithelial cells, bound to one another by tight junctions, undergo apoptosis (cellular implosion) and draw adjacent cells toward the site of cell loss. It is proposed that this resultant “apoptotic force” would serve to pull the inferior follicle upward in catagen.

by the connective tissue sheath and its glassy membrane.46 Recently, smooth muscle α-actin was identified in the cells of the connective tissue sheath.95 Another possibility, not yet proven experimentally, is that there is an apoptotic force (Fig. 7.3). This force would be generated in an epithelial structure where cell contacts are tight and where cells are not dividing. In this case, when individual cells implode from the apoptotic process the epithelial sheet would necessarily contract due to the tight cell contacts. An upward vectorial force would result, pulling the lower follicle up. What the initiating signal for catagen might be is as obscure as the signal for anagen induction. In the laboratory, catagen can be induced by dexamethasone and this effect can be abrogated by cyclosporine A.96 Apoptotic-like changes can also be induced in follicles by EGF97 and TGF-β2.88 Recently, it was found serendipitously that fibroblast growth factor-5 plays some role in catagen induction.98,99 In the FGF-5 null mouse, for both the transgenic mouse and the spontaneously found mutant (Angora), anagen is prolonged (catagen is delayed) and the resultant hair shafts are longer. It is notable that eventually, even in the absence of this growth factor, catagen is induced, indicating that FGF-5 is not the only controlling molecule. In catagen major changes occur in the mesenchymal component of the follicle as well. The proteoglycan character of the papilla changes as the follicle enters catagen, e.g., versican expression drops100 and protein elements such as nexin-1101 and alkaline phosphatase change in concentration.46,49 Recently, it has been found that some genes are activated during catagen, such as osteopontin.102

Growth of the Hair Follicle

121

Telogen Telogen is defined as the phase during which hair follicle growth, and thus hair shaft growth, comes to a halt. This is manifest by the lack of cell division in the telogen germ. Although it is generally agreed that no cell growth occurs during this phase, that telogen is an inactive phase has not yet been conclusively demonstrated. Since we do not know how follicle growth is controlled, we cannot exclude the possibility that telogen actively suppresses hair growth. A paradigm for the latter explanation is the bristle appendage of Drosophila integument,103 which is controlled by a suppressive signal of the Achaete-scute gene complex. As to that, the possibility that telogen skin contains an anagen inhibitor60 could support that mechanism. If the cycle is controlled by a negative telogen-associated component, then the telogen follicle might play an active role in follicle cycle control.

Exogen Once the growth cycle is complete, including the phase of telogen, the hair shaft will be shed. This process occurs in that part of the follicle independent of the new germ and its growth. There is no doubt, from a clinical and comparative biological respect, that shedding is exquisitely controlled and that it occurs independent of the new cycle. In fact, it is most common in mammals to regrow a new hair shaft before the resting shaft sheds; this assures that the animal is never naked. Since shedding is conceived by us and others in the field as a separate and distinctive process of the hair cycle, we propose to name it exogen (Lt. ex, out of; Lt. generare, give rise to) because this word is etymologically consistent with the phase names suggested by Dry.20 It is notable that in that paper, Dry20 dedicated as much attention to the shedding phase as he did to the other phases though he did not name it. Other workers have also discussed shedding as a phase independent of the other phases.52,105,106 Exogen is seen as the phase of the cycle wherein the signals for shaft release are given and the processes for that release are initiated and successfully executed. Exogen ends when the shaft is liberated. That the exogen phase of one cycle is independent of the growth or exogen phases of subsequent cycles is clear from the condition of trichostasis, physiological or pathological, in which multiple new shafts are formed from the same follicle and are retained within that follicle. Because of this relationship, it is almost certain that shedding involves much more than mere mechanical trauma. and that follicle growth and shedding are independent events. As mentioned above, antler shedding and growth initiation also appear to be independent events.16 From a clinical standpoint the critical phase of the cycle for a patient suffering from hair loss is exogen: Despite the growth phase, if exogen were prevented, there would be no presenting complaint. It is important to recognize that the onset of exogen may vary with the season,105 with follicle type20 and with follicle location.41 In animals whose pelage grows synchronously, exogen occurs at certain times of the year. There is an implicit assumption that the induction of telogen relates to the induction of exogen. That may or may not be true when considering the marked variation of exogen start in animals (e.g., the delayed exogen of the mink over the winter). In this regard Headington64 identified different types of telogen hair shedding (exogens) associated with different clinical conditions, although the exogen phase in all cases was probably the same. The parameters controlling exogen, the shedding of hair and wool, are for the most part ill-defined. For sheep its control is considered to be genetic since the pattern of wool loss is dependent on the breed.15 Seasonal molting is controlled by endocrine messages under the coordinating influence of the pineal gland and environmental signals. Pinealectomy, or agents which prevent a drop in prolactin levels, abrogate seasonal molting.27 As noted

122

Molecular Basis of Epithelial Appendage Morphogenesis

above, circulating prolactin levels correlate inversely with melatonin levels, up during the summer and down during the winter months. Despite its obvious importance, neither the morphological nor the biochemical mechanisms of exogen have been explored. We recognize that the telogen club hair rests as a keratin brush adherent to the telogen outer root sheath. The next morphological alteration we are certain of is that a club hair is shed: The shed shaft has a bulbous keratinized root and the parent follicle manifests an indentation in the wall of the deep pilary canal. A priori, it would appear that the release mechanism would involve an enzymatic action, but which enzymes, from which cells and under what controls are completely unknown.

Hair Follicle Growth Controls—A Mechanistic Overview Many molecules are found to be expressed over the growth cycle, some of which are characteristic of certain phases.6 The significance of any of these molecules, besides the structural ones, is not yet clear.

Hormones The influence of hormones on hair growth has been reviewed.107,108 With regard to the wool growth cycle, there are at least three influential endocrine glands: the anterior pituitary, thyroid and adrenal cortex. Hormones are believed to play one of three roles: 1. a permissive role, necessary for normal hair growth but without an effect on the rate of growth; 2. a regulatory role, in which variation in secretion rate affects the production and, perhaps, the shedding of hair; and, finally, 3. those molecules that have an effect on hair growth but only at pharmacological levels.109 In general, hormones of the pituitary and thyroid gland stimulate hair growth. Corticosteroids at high concentrations suppress hair growth and at low concentrations may be stimulatory. Although the inhibitory action of corticosteroids has been shown to act on the initiation of a growth wave, once hair growth has started the treatment has no effect.65,110 It is notable that dexamethasone has been reported to initiate catagen.96 Thyroid and adrenal corticoid hormones stimulate the molt and loss of club hairs.105 Androgens stimulate specialized follicles but will inhibit growth of follicles (e.g., scalp follicles) in the appropriate host. Their important role in the development of some follicles is well documented.111 In the vole,112 androgens inhibit the growth of pelage hairs while, conversely, removal of the testes (with reduction of systemic androgens) stimulates hair growth. Evidence for a role of androgens in control of the cycle per se has not been presented; however, that these hormones impact the epidermis (e.g., ref. 113) and the sebaceous gland114 may generate an indirect effect on the cycle. It is notable that androgens will have an effect on the type of shaft and follicle produced in androgen sensitive regions. Although androgens influence the expression of growth factors that stimulate follicle growth in vitro (e.g., FGF,115 KGF,116 IGF117), how and if androgens influence the cycle are less clear at this time. A new deer model studying this follicle size circannual switch has been developed.118,119 Work suggesting a role of estrogen in hair cycle control has been put forth but not confirmed.120

Inflammatory and Immune Elements The importance of the immune system in the hair cycle was first intimated by the finding that the lower cycling portion of the follicle is immunologically unique.89 In anagen, there is an absence of class I MHC expression on most cells of the lower follicle and a presence of chondroitin proteoglycan within the follicle sheath and papilla. Catagen initiation is

Growth of the Hair Follicle

123

associated with the reexpression of class I MHC antigen on all follicle epithelium; concomitantly chrondroitin sulfate proteoglycan disappears from the sheath and papilla.90,121 This inverse relationship has been thought to play a role in modulating access of immune cells to the follicle. In catagen a large number of activated macrophages aggregate around the follicles.122 During the entire hair cycle, Vgamma 3-TCR bearing lymphocytes are found in the epidermis, the bulb and the bulge areas. During early anagen the number of γδ T cells increase in the interfollicular epidermis and the suprainfundibular portion of the hair bulb. There is then a cycle dependent proliferation of γδ T cells in mouse skin.123 While the studies indicate that MHC class I antigens are absent in the matrix and inner root sheath of growing hair follicles, nonclassical MHC class I antigens, showing strong Qa-2 immunoreactivity, are found in the distal hair follicle during the entire hair cycle. This region is restricted to the area populated with γδ T cells.124 Other implications that the immune system plays a role in the hair cycle include the expression of immune mediators during the cycle,35 the fact that immune mediators affect hair growth in vitro,125 and that potent immunosuppressive drugs, Cyclosporine A and FK 506, stimulate hair growth,66,126-129 in addition to the immune marker changes of the lower follicle during anagen and the inflammatory infiltrate that accumulates about the follicle in catagen. The significant association of follicular growth and immunological abnormalities in the nude mouse has implicated the whn gene in the functioning of both systems.130-132 Mast cells are observed to degranulate during anagen, so that by late anagen there is a radical decrease in mast cells, histamine and heparin.104,133 The greatest mast cell degranulation occurs in early anagen and the lowest number of degranulated mast cells are found in catagen.134 Because the number and granulation status of mast cells change during the murine hair cycle, it is thought that these cells play a growth modulatory function.135 The impression is that mast cells play some role in anagen development.

Neural The neural influence on the hair cycle is another aspect of hair biology that needs careful study. It has long been appreciated that the follicle is richly innervated, but not that the innervation changes with the cycle.136 This observation, as well as the report that hair follicle growth is induced by substance P,137 implicates an important neural component.138

Vascular As it is a rapidly dividing tissue, it is no surprise that the growing hair follicle is richly supplied with vessels. The growing follicle is surrounded by a network of vessels that arise from the deep dermal plexus.139-141 For larger follicles the papilla receives its own vascular supply. With the onset of catagen, the vascular net collapses about the regressing follicle and apparently remains in a quiescent state until the initiation of the next cycle. Because follicular atrophy begins before vascular collapse, it is thought that the signal for catagen is not from the vessels. That few new vessels are formed over the cycle was supported by kinetic studies that showed greater thymidine uptake in vessels surrounding anagen growth.142 Angiogenic stimuli within the hair follicle have been reported to be within the follicular epithelium, 143 but which of the known angiogenic growth factors that have been identified within the epithelium (VEGF, PDGF, EGF, TGF-β) are responsible for that role is not yet apparent. The current impression is that the dermal vessels respond to the hair cycle and support it, but they do not provide the initial stimulus to growth. The vascular response, then, is part of the cross-talk between these two tissues, the epithelium and the mesenchyme.

124

Molecular Basis of Epithelial Appendage Morphogenesis

Conclusions A unique developmental organ with parallels to the antler, the endometrium and the feather, hair grows through cycles with a long period of from months to years. The cycle is influenced by genetics, and by hormones that are themselves often mediated by environmental influences. Our knowledge of the cycle is phenomenological but certain factors would appear to be important to its regulation. The factors include the immune system, neural factors, vascular factors, cytokines, growth factors, and epithelial mesenchymal interactions. In his classical review, Chase46 concluded his discussion with a list of the unsolved problems in hair biology of his day. His major concern was with the control of the cycle itself. Although we have more experimental data, we do not have much more insight today. But, optimistically, we have powerful tools that he did not have, particularly with regard to animal models, tissue culture techniques, means of identifying and isolating the relevant genes and, in the immediate future, knowing all the molecules relevant to hair growth. Obviously only by integrating those components will we meet the challenge of Chase.

Acknowledgments The authors are indebted to Prof. Ralf Paus for discussion and the opportunity to review manuscripts in press.

References 1. Kondo T, Strayer CA, Kulkarni R et al. Circadian rhythms in prokaryotes: Luciferase as a reporter of circadian gene expression in cyanobacteria. Proc Natl Acad Sci USA 1993; 90:5672-5676. 2. Smaaland R. Circadian rhythm of cell division. In: Progress in Cell Cycle Research. Meijer L, Guidet S, Vogel L, eds. NY: Plenum Press, 1996:241-266. 3. Rubin N, Scheving LE. Circadian rhythm. J Invest Dermatol 1983; 80:79-80. 4. Scheving LE, Tsai TH, Scheving LA. Chronobiology of the intestinal tract of the mouse. Am J Anat 1983; 168:433-465. 5. Tsai TH, Scheving LE, Marques N et al. Circadian-infraradian intermodulation of corneal epithelial mitoses in adult female rats. Prog Clin Biol Res 1987; 227A:93-198. 6. Stenn KS, Combates NJ, Eilertsen KJ et al. Hair follicle growth controls. Dermatol Clin 1996; 14:543-558. 7. Paus R. Control of the hair cycle and hair disease as cycling disorders. Curr Opin Dermatol 1996a; 3:248-258. 8. Goss RJ. Wound healing and antler regeneration. In: Epidermal Wound Healing. Maibach HI, Rovee DT, eds. Year Book Med Publ Chicago, 1972:219-228. 9. Brugureolle B, Lemmer B. Recent advances in chronopharmacokinetic: Methodological problems. Life Sci 1993; 52:1809-1824. 10. Takahashi JS. Circadian-clock regulation of gene expression. Curr Opin Gen Dev 1993; 3:301-309. 11. Hastings M. What makes the clock tick? Cur Biol 1994; 4:720-723. 12. Dunlap JC. Genetic and molecular analysis of circadian rhythms. Ann Rev Genet 1996; 30:579-601. 13. King DP, Zhao Y, Sangoram AM et al. Positional cloning of the mouse circadian Clock gene. Cell 1997; 89:641-653. 14. Kay SA. PAS, present, and future: Clues to the origins of circadian clocks. Science 1997; 276:753-754. 15. Ryder ML, Stephenson SK. Seasonal changes in the fleece and their hormonal control. Chap 12 In: Wool Growth, London: Acad Press, 1968:593-625. 16. Goss, RJ. Deer Antlers, Regeneration, Function, and Evolution. NY: Acad Press, 1983:1-316. 17. Ying SY. Inhibins, activins, and follistatins: Gonadal proteins modulating the secretion of follicle-stimulating hormone. Endocr Rev 1988; 9:267-293.

Growth of the Hair Follicle

125

18. Borgens RB. Are limb development and limb regeneration both initiated by an integumentary wounding? A hypothesis. Differentiation. 1984; 28:87-93. 19. Ralph MR, Hurd MW. Circadian pacemakers in vertebrates. Ciba Found Sympos 1995; 183:67-87. 20. Dry FW. The coat of the mouse (Mus musculus). J Genet 1926; 16:287-340. 21. Bassett CF, Llewellyn LM. The molting and fur growth pattern in the adult mink. Amer Midl Nat 1949; 42:751-756. 22. Parry AL, Nixon AJ, Craven AJ et al. The microanatomy, cell replication, and keratin gene expression of hair follicles during a photoperiod-induced growth cycle in sheep. Acta Anat 1995; 154:283-299. 23. Foldes A, Hoskinson RM, Baker P et al. Effect of immunization against melatonin on seasonal fleece growth in feral goats. J Pineal Res 1992; 13:85-94. 24. Slominski A, Chassalevris N, Mazurkiewica J et al. Murine skin as a target for melatonin bioregulation. Exp Dermatol 1994; 3:45-50. 25. Smale L, Nelson RJ, Zucker I. Daylength influences pelage and plasma prolactin concentrations but not reproduction in the prairie vole, Microtus ochrogaster. J Reprod Fert 1988; 83:99-106. 26. Rose J, Garwood T, Jaber B. Prolactin receptor concentrations in the skin of mink during the winter fur growth cycle. J Exp Zool 1995; 271:205-210. 27. Martinet L, Allain D, Weiner C. Role of prolactin in photoperiodic control of molting in the mink (Mustela vison). J Endocr 1984; 103:9-15. 28. Clarke LA, Edery M, Loudon AS et al. Expression of the prolactin receptor gene during the breeding and non-breeding season in red deer (Cervus elaphus): Evidence for the expression of two forms in the testis. J Endocrinol 1995; 146:313-321. 29. Ebling FJ, Johnson E. Hair growth and its relation to vascular supply in rotated skin grafts and transposed flaps in the albino rat. J Embryol Exp Morph 1959; 7:417-430. 30. Ebling FJ, Hervey GR The activity of hair follicles in parabiotic rats. J Embryol Exp Morph 1964; 12:425-438. 31. Johnson E. Quantitative studies of hair growth in the albino rat. 2. The effect of sex hormones. J Endocr 1958; 16:351-359. 32. Durward A, Rudall KM. Hair growth in the rat. J Anat 1949; 83:325-335. 33. Slee J, Carter HB. Fibre shedding and fibre follicle relationships in the fleeces of Wiltshire Horn x Scottish Blackface sheep crosses. J Agric Sci, Camb 1962; 58:309-326. 34. Stenn KS, Prouty SM, Seiberg M. Molecules of the cycling hair follicle—a tabulated review. J Dermatol Sci 1994; 7(Suppl) S109-S124. 35. Seiberg M, Marthinuss J, Stenn KS. Changes in the expression of apoptosis-associated genes in skin mark early catagen. J Invest Dermatol 1995; 104:78-82. 36. Raju U, Koumenis C, Nunez-Reguiero M et al. Alteration of the phase and period of a circadian oscillator by a reversible transcription inhibitor. Science 1991; 253:673-675. 37. Vitaterna MH, King DP, Chang A-M et al. Mutagenesis and mapping of a mouse gene, clock, essential for circadian behavior. Science 1994; 264:719-725. 38. Holbrook KS, Fisher C, Dale BA et al. Morphogenesis of the hair follicle during the ontogeny of human skin. Chap 2 In: The Biology of Wool and Hair. GE Rogers, PJ Reis, KA Ward, Marshall RC, eds London: Chapman & Hall 1989:15-35. 39. Pecoraro V, Astore I, Barman JM. Cycle of the scalp hair of the newborn child. J Invest Dermatol 1964; 43:145-147. 40. Barman JM, Pecoraro V, Astore I et al. The first stage in the natural history of the human scalp hair cycle. J Invest Dermatol 1967; 48:138-142. 41. Saitoh M, Uzuka M, Sakamoto M. Human hair cycle. J Invest Dermatol 1970; 54:65-81. 42. Pecoraro V, Astore I, Barman JM. Growth rate and hair density of the human axilla. J Invest Dermatol 1971; 56:362-365. 43. Seago SV, Ebling FJG. The hair cycle on the human thigh and upper arm. Brit J Dermatol 1985; 113:9-16. 44. Orentreich N. Scalp hair replacement in man. Adv Skin Biol 1969; 9:99-108.

126

Molecular Basis of Epithelial Appendage Morphogenesis

45. Randall VA, Ebling FJG. Seasonal changes in human hair growth. Brit J Dermatol 1991; 124:146-151. 46. Chase HB. Growth of the hair. Physiol Rev 1954; 34:113-126. 47. Chase HB, Montagna W, Malone JD. Changes in the skin in relation to the hair growth cycle. Anat Rec 1953; 116:75-82. 48. Hansen LS, Coggle JE, Wells J et al. The influence of the hair cycle on the thickness of mouse skin. Anat Rec 1984; 210:569-573. 49. Handjiski BK, Eichmuller S, Hofmann U et al. Alkaline phosphatase activity and localization during the murine hair cycle. Brit J Dermatol 1994; 131:303-310. 50. Hofmann U, Tokura Y, Nishijima T et al. Hair cycle dependent changes in skin immune functions: anagen associated depression of sensitization for contact hypersensitivity in mice. J Invest Dermatol 1996; 106:598-694. 51. Hardy MH. The secret life of the hair follicle. Trends in Genet 1992; 8:55-61. 52. Straile WE, Chase HB, Arsenault C. Growth and differentiation of hair follicles between periods of activity and quiescence. J Exp Zool 1961; 148:205-216. 53. Argyris TS. The effects of wounds on new tissue in an adult mammal. Symp Soc Exp Biol. 1956; 11:235-254. 54. Argyris TS. The growth promoting effects of wounds on hair follicles already stimulated by plucking. Anat. Rec 1962; 143:183-188. 55. Argyris TS, Argyris BF. Factors affecting the stimulation of hair growth during wound healing. Anat Rec 1962; 142:139-145. 56. Silver AF, Chase HB, Arsenault CT. Early anagen initiated by plucking compared with early spontaneous anagen. Adv Biol Skin 1969; 9:265-286. 57. Chase HB, Eaton GJ. The growth of hair follicles in waves. Ann NY Acad Sci 1959; 83:365-368. 58. Chase HB. The physiology and histochemistry of hair growth. J Soc Cosmet Chem 1955; 6:9-14. 59. Bullough WS, Laurence EB. The control of epidermal mitotic activity in the mouse. Proc Roy Soc B 1960; 151:517-536. 60. Paus R, Stenn KS, Link RE. Telogen skin contains an inhibitor of hair growth. Brit J Dermatol 1990; 122:777-784. 61. Johnson E, Ebling FJ. The effect of plucking hairs during different phases of the follicular cycle. J Embryol Exp Morph 1964; 12:465-474. 62. Lynfield YL. Effect of pregnancy on the human hair cycle. J Invest Dermatol 1960; 35:323-327. 63. Messenger AG. The control of hair growth: An overview. J Invest Dermatol 1993; 101:4s-9s. 64. Headington JT. Telogen effluvium new concepts and review. Arch Dermatol 1993; 129:356-363. 65. Stenn KS, Paus R, Dutton T et al. Glucocorticoid effect on hair growth initiation: A reconsideration. Skin Pharm 1993; 6:125-134. 66. Paus R, Stenn KS, Link RE Jr. The induction of anagen hair follicle growth in telogen mouse skin by cyclosporin A administration. Lab Invest 1989; 60:365-369. 67. Yamamoto S, Jiang H, Kato R. Stimulation of hair growth by topical application of FK506, a potent immunosuppressive agent. J Invest Dermatol 1994; 102:160-164. 68. Buhl AE, Waldon DJ, Miller BF et al. Differences in activity of minoxidil and cyclosporin A on hair growth in nude and normal mice. Lab Invest 1990; 62:104-107. 69. Paus R, Heinzelmann T, Schultz KD et al. Hair growth induction by substance P. Lab Invest 1994f; 71:134-140. 70. Shimaoka A, Tsuboi R, Jindo T et al. Hepatocyte growth factor/scatter factor expressed in follicular papilla cells stimulates hair growth in vitro. J Cell Physiol 1995; 165:333-338. 71. Danilenko DM, Ring BD, Yanagihara D et al. Keratinocyte growth factor is an important endogenous mediator of hair follicle growth, development and differentiation. Normalization of the nu\nu follicular differentiation defect and amelioration of chemotherapy-induced alopecia. Amer J Pathol 1995; 147:145-154.

Growth of the Hair Follicle

127

72. Cotsarelis G, Sun T-T, Lavker RM. Label-retaining cells reside in the bulge area of pilosebaceous unit: Implications for follicular stems cells, hair cycle and skin carcinogenesis. Cell 1990; 61:1329-1337. 73. Reynolds AJ, Jahoda CAB. Inductive properties of hair follicle cells. Ann NY Acad Sci 1991; 642:226-241. 74. Kobayashi K, Rochat A, Barrandon Y. Segregation of keratinocyte colony-forming cells in the bulge of the rat vibrissa. Proc Natl Acad Sci 1993; 90:7391-7395. 75. Rochat A, Kobayashi K, Barrandon Y. Location of stem cells of human hair follicles by clonal analysis. Cell 1994; 76:1063-1073. 76. Yang J-S, Lavker RM, Sun T-T. Upper human hair follicle contains a subpopulation of keratinocytes with superior in vitro proliferative potential. J Invest Dermatol 1993; 101:652-659. 77. Wilson CL, Sun TT, Lavker RM. Cells in the bulge of the mouse telogen follicle give rise to the lower anagen follicle. Skin Pharmacol 1994; 7:8-11. 78. Weinberg WC, Brown PD, Stetler-Stevenson WG et al. Growth factors specifically alter hair follicle cell proliferation and collagenolytic activity alone or in combination. Differentiation 1990; 45:168-178. 79. Paus R, Krejci-Papa N, Li L et al. Correlation of proteolytic activities of organ cultured intact mouse skin with defined hair cycle stages. J Dermatol Sci 1994e; 7:202-209. 80. Williams DD, Stenn KS. Transection level dictates the pattern of hair follicle sheath growth in vitro. Dev Biol 1994; 165:469-479. 81. Couchman JR, Gibson WT. Expression of basement membrane components through morphological changes in the hair growth cycle. Dev Biol 1985; 108:290-298. 82. Jahoda CAB, Mauger A, Bard S et al. Changes in fibronectin, laminin and type IV collagen distribution relate to basement membrane restructuring during the rat vibrissa follicle hair growth cycle. J Anat 1992; 181:47-60. 83. Wessells NK, Roessner KD. Nonproliferation in dermal condensations of mouse vibrissae and pelage hairs. Dev Biol 1965; 12:419-433. 84. Silver AF, Chase HB. DNA synthesis in the adult hair germ during dormancy (telogen) and activation (early anagen). Dev Biol 1970; 21:440-451. 85. Link RE, Paus R, Stenn KS et al. Epithelial growth in cultured rat vibrissae follicles requires mesenchymal contact via native extracellular matrix. J Invest Dermatol 1990; 95:202-207. 86. Birbeck MSC, Mercer EH. The electron microscopy of the human hair follicle. Part 1. Introduction and the hair cortex. J Biophysic and Biochem Cytol 1957a; 3:203-213. 87. Birbeck MSC, Mercer EH. The electron microscopy of the human hair follicle. Part 2. The hair cuticle. J Biophysic and Biochem Cytol 1957b; 3:215-221. 88. Soma T, Ogo M, Suzuki J et al. Analysis of apoptotic cell death in human hair follicle. J Invest Dermatol 1997; 108:581abstr. 89. Harris TJ, Ruiter DJ, Mihm MC et al. Distribution of major histocompatibility antigens in normal skin. Brit J Dermatol 1983; 109:623-633. 90. Westgate GE, Craggs RI, Gibson WT. Immune privilege in hair growth. J Invest Dermatol 1991a; 97:417-420. 91. DeWeert J, Kint A, Geert ML. Morphological changes in the proximal area of the rat’s hair follicle during early catagen. Dermatol Res 1982; 272:79-92. 92. Weedon D, Strutton G. The recognition of the early stages of catagen. Amer J Dermatopath 1984; 6:553-555. 93. Lindner G, Bochkarev VA, Botchkareva N et al. Towards generation of an “apoptomap” of the murine hair follicle. J Invest Dermatol 1997; 108:620 abstr. 94. Kligman AM. The human hair cycles. J Invest Dermatol 1959; 33:307-316. 95. Reynolds AJ, Chaponnier C, Jahoda CA et al. A quantitative study of the differential expression of alpha-smooth muscle actin in cell populations of follicular and non-follicular origin. J Invest Dermatol 1993; 101:577-583.

128

Molecular Basis of Epithelial Appendage Morphogenesis

96. Paus R, Handjiski B, Czarnetzki BM et al. A murine model for inducing and manipulating hair follicle regression (catagen): Effects of dexamethasone and cyclosporin A. J Invest Dermatol 1994a; 103:143-147. 97. Philpott MP, Kealey T. Effect of EGF on the morphology and patterns of DNA synthesis in isolated human hair follicles. J Invest Dermatol 1994; 102:186-191. 98. Hebert JM, Rosenquist T, Gotz J et al. FGF5 as a regulator of the hair growth cycle: Evidence from targeted and spontaneous mutations. Cell 1994; 78:1017-1025. 99. Sundberg JP, Rourk MH, Boggess D et al. Angora mouse mutation: Altered hair cycle, follicular dystrophy, phenotypic maintenance of skin grafts, and changes in keratin expression. Vet Pathol 34: (in press). 100. DuCros DL, Le Baron RG, Couchman JR. Association of versican with dermal matrices and its potential role in hair follicle development and cycling. J Invest Dermatol 1995; 105:426-431. 101. Yu D-W, Yang T, Sonoda T et al. Message of nexin 1, a serine protease inhibitor, is accumulated in the follicular papilla during anagen of the hair cycle. J Cell Sci 1995; 108:3867-3874. 102. Yu D-W, Yang T, Sonoda T et al. Osteopontin gene is upregulated during the catagen phase of the hair cycle. J Invest Dermatol 1997; 108:653abstr. 103. Fristrom D, Fristrom JW. The metamorphic development of the adult epidermis. Chap 14 In The Development of Drosophila melanogaster. Bate M, Arias AM, eds. Cold Spring Harbor Laboratory Press Vol II 1993:843-897. 104. Paus R, Maurer M, Slominski AM et al. Mast cell involvement in murine hair growth. Dev Biol 1994d; 163:230-240. 105. Johnson E. Molting cycles. Mammalian Rev. 1972; 1:198-208. 106. Pinkus H. Factors in the formation of club hair. Chap 9 In: Hair, Trace Elements and Human Illness. Brown AC, Crounse RG, eds. NY: Praeger Scientific Press, 1980:147-154. 107. Ebling FJG, Hale PA, Randall VA. Hormones and hair growth. In: Goldsmith LA, ed. Physiology, Biochemistry, and Molecular Biology of the Skin, Second Edition. New York: Oxford Univ Press, 1991:660-696. 108. Randall VA. Androgens and human hair growth. Clin Endocrin 1994; 40:439-457. 109. Ferguson KA, Wallace ALC, Lindner HR Hormonal regulation of wool growth. In: Biology of the Skin and Hair Growth, Lyne AG, Short BF eds. Sydney: Angus and Robertson, 1965:655-677. 110. Mohn MP. The effect of different hormonal states on the growth of hair in rats. In: The Biology of Hair Growth. Montagna W,Ellis RA, eds.NY: Academic Press, 1958:336- 399. 111. Rosenfield RL, Deplewski D. Role of androgens in the developmental biology of the pilosebaceous unit. Amer J Med 1995; 98:80S-88S. 112. Khateeb AA, Johnson E. Seasonal changes of pelage in the vole (Microtus agrestis) 1. Correlation with changes in the endocrine glands. Gen Comp Endocr 1971; 16:217-238. 113. Hanley K, Rassner U, Jiang Y et al. Hormonal basis for the gender difference in epidermal barrier formation in the fetal rat. Acceleration by estrogen and delay by testosterone. J Clin Invest 1996; 97:2576-2584. 114. Imperato-McGinley J, Gautier T, Cai L-Q et al. The androgen control of sebum production. Studies of subjects with dihydrotestosterone deficiency and complete androgen insensitivity. J Clin Endocrinol Metab 1993; 76:524-528. 115. Ashton WS, Degnan BM, Daniel A et al. Testosterone increases insulin-like growth factor1and insulin-like growth factor-binding protein. Ann Clin Lab Sci 1995; 25:381-388. 116. Fasciana C, van der Made AC, Faber PW et al. Androgen regulation of the rat keratinocyte growth factor (KGF/FGF7) promoter. Biochem Biophys Res Commun 1996; 220:858-863. 117. Crawford BA, Handelsman DJ. Androgens regulate circulating levels of insulin-like growth factor IGF-1 and IGF binding protein-3 during puberty in male baboons. J Clin Endocrinol Metab 1996; 81:65-72. 118. Heydon MJ, Milne JA, Brindlow BR et al. Manipulating melatonin in red deer (Cervus elaphus): Differences in the response to food restriction and lactation on the timing of the breeding season and prolactin-dependent pelage changes. J Exp Zool 1995; 273:12-20.

Growth of the Hair Follicle

129

119. Thornton MJ, Kato S, Hibbers NA et al. Ability to culture dermal papilla cells from red deer (Cervus elaphus) hair follicles with differing hormonal responses in vivo offers a new model for studying the control of hair follicle biology. J Exp Zool 1996; 275:452-458. 120. Oh H-S, Smart RC. An estrogen receptor pathway regulates the telogen-anagen hair follicle transition and influences epidermal cell proliferation. Proc Natl Acad Sci 1996; 93:12525-12530. 121. Westgate GE, Messenger AG, Watson LP et al. Distribution of proteoglycans during the hair growth cycle in human skin. J Invest Dermatol 1991b; 196:191-195. 122. Gibson WT, Westgate GE, Craggs RI. Immunology of the hair follicle. Ann NY Acad Sci 1991; 642:291-300. 123. Paus R, Hofmann U, Eichmuller S et al. Distribution and changing density of gammadelta T cells in murine skin during the induced hair cycle. Brit J Dermatol 1994b; 130:281-289. 124. Paus R, Eichmuller S, Hofmann U et al. Expression of classical and non-classical MHC class I antigens in murine hair follicles. Brit J Dermatol 1994c; 131:177-183. 125. Philpott MP, Sanders DA, Bowen J et al. Effects of interleukins, colony-stimulating factor and tumour necrosis factor on human hair follicle growth in vitro: A possible role for interleukin-1 and tumour necrosis factor-α in alopecia areata. Brit J Dermatol 1996; 135:942-948. 126. Pendry A, Alexander P. Stimulation of hair growth on nude mice by cyclosporin A. In: White DJG, ed. Cyclosporin A. Elsevier Biomedical Press, 1982:77. 127. Sawada M, Terada N, Taniguchi H et al. Cyclosporin A stimulates hair growth in nude mice. Lab Invest 1987; 56:684-686. 128. Jiang H, Yamamoto S, Kato R. Induction of anagen in telogen mouse skin by topical application of FK506, a potent immunosuppressant. J Invest Dermatol 1995; 104:523-525. 129. Iwabuchi T, Maruyama T, Sei Y et al. Effects of immunosuppressive peptidyl-prolyl cistrans isomerase (PPIase) inhibitors, cyclosporin A, FK 506, ascomycin and rapamycin, on hair growth initiation in mouse: Immunosuppression is not required for new hair growth. J Dermatol Sci 1995; 9:64-69. 130. Nehls M, Pfeifer D, Schorpp M et al. New member of the winged-helix protein family disrupted in mouse and rat nude mutations. Nature 1994; 372:103-107. 131. Segre J, Nemhauser J, Taylor B et al. Positional cloning of the nude Locus: Genetic, physical, and transcription maps of the region and mutations in the mouse and rat. Genomics 1995; 28:387-398. 132. Brissette JL, Li J, Kamimura J et al. The product of the mouse nude locus, whn, regulates the balance between epithelial cell growth and differentiation. Gene Dev 1996; 10:2212-2221. 133. Moretti GM, Ranabou EM, Rebora A. The hair cycle re-evaluated. Int J Dermatol 1976; 15:2777-2781. 134. Paus R, Maurer M, Slominski A et al. Mast cell involvement in murine hair growth. Dev Biol 1994g; 163:230-240. 135. Botchkarev VA, Paus R, Czarnetzki BM et al. Hair cycle-dependent changes in mast cell histochemistry in murine skin. Arch Dermatol Res 1995; 287:683-686. 136. Botchkarev VA, Eichmuller S, Johansson O et al. Hair cycle-dependent plasticity of skin and hair follicle innervation in normal murine skin. J Comp Neurol (in press). 137. Paus R, Heinzelmann T, Schultz KD et al. Hair growth induction by substance P. Lab Invest 1994h; 71:134-140. 138. Paus R, Peters EMJ, Eichmuller S et al. Neural mechanisms of hair growth control. J Invest Dermatol (in press). 139. Montagna W, Ellis RA. Histology and cytochemistry of human skin. XIII. The blood supply of the hair follicle. J Nat Cancer Inst 1957; 19:451-463. 140. Durward A, Rudall KM. The vascularity and patterns of growth of hair follicles. In: Montagna W, Ellis RA, eds. The Biology of Hair Growth. New York: Acad Press, 1958:189-218. 141. Ellis RA. Vascular patterns associated with catagen hair follicles in the human scalp. Ann NY Acad Sci 1959; 83:448-457.

130

Molecular Basis of Epithelial Appendage Morphogenesis

142. Sholley MM, Cotran RS. Endothelial DNA synthesis in the microvasculature of rat skin during the hair growth cycle. Amer J Anat 1976; 147:243-254. 143. Stenn KS, Fernandez LA, Tirrell SJ The angiogenic properties of the rat vibrissa hair follicle associate with the bulb. J Invest Dermatol 1988; 90:409-411. 144. Montagna W, Parakkel PF. The Structure and Function of Skin. New York: Acad Press, 1974: 250. 145. Tsonis PA. Limb regeneration. Cambridge: Cambridge University Press, 1996: 1-201.

CHAPTER 8

Development of Melanocytes from Neural Crest Progenitors Laure Lecoin, Ronit Lahav, Elisabeth Dupin and Nicole Le Douarin

Introduction

T

he pigmented cells of the body, the melanocytes, have long been a favored cell type for studies of the genetic and epigenetic factors involved in cell differentiation, partly because pigment makes these cells easily recognizable without any staining, but also because of the astonishing variety of the patterns (spots, zebra, pictures...) due to the distribution of pigment cells in the skin of animals. The fact that pigment cell precursors (melanoblasts) can be successfully transplanted to new tissue environments has made possible experimental analysis of the relative roles of genotype and environment in directing melanocyte differentiation in many different species like amphibians, avians and mice. The wealth of different genotypes affecting coat color in the mouse also enabled the identification of different loci that act either through the cellular environment or within the melanoblasts to influence their development and function. Recently, with advances in the field of molecular biology, the gene products of some loci have been identified. Thus, genetic and environmental influences could be attributed to isolated factors, which in turn enabled a detailed characterization of the nature of these influences in the process of melanocyte differentiation. The first part of this chapter will summarize the general knowledge about neural crestderived pigment cell development. The second part will aim to describe how melanoblast distribution is controlled and may account for the pigmentation patterns observed on the skin.

Early Migration and Commitment of Pigment Cells The Neural Crest, Source of Pigment Cells in Vertebrates The neural crest origin of pigment cells has been demonstrated by transplantation and culture experiments of different tissues isolated from amphibian,1,2 avian3,4 and mouse5 embryos. The neural crest is a transient and pluripotent cell population that develops at the time of closure of the neural tube. This cell population undergoes an epithelio-mesenchymal transition and neural crest cells start migrating from the dorsal part of the neural tube until they reach their final locations of differentiation. The derivatives of the neural crest include neurons and glial cells of the peripheral nervous system, the majority of the head mesenchyme, some endocrine and paraendocrine cells and melanocytes (see ref. 6 for a review). All the pigment cells of the body originate from the neural crest except the pigmented cells of the retina, which derive from the neural epithelium of the optic vesicle. Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

132

Molecular Basis of Epithelial Appendage Morphogenesis

Migration Pathways of Pigment Cell Precursors The neural crest migration pathways have been mainly studied in the avian embryo because of its accessibility. So we will first describe migration of melanoblasts in the avian embryo before mentioning the main differences observed in the other species. Pioneer studies7-10 established the general timing of melanoblast migration in the avian embryo by examining the ability to generate pigment cells of embryonic regions explanted at different developmental stages. The lateral part of the somite is invaded by melanoblasts at stage 33 somites; wing buds begin to be colonized by these cells at embryonic day 3.5 (E3.5) and leg buds at E4.5. However, at these early stages, melanoblasts cannot be distinguished from the surrounding mesenchymal cells, since pigment production begins later in development. The debate about whether pigment cell precursors migrate within the ectoderm or in the underlying mesenchyme was solved by the use of the quail-chick marker system. This stable and inheritable marker is based on the morphological difference that exists, during the interphase, between the chick cell nuclei and those of the Japanese quail.11 The method consists in replacing a fragment of chick neural primordium by its counterpart from a quail neural primordium at the same developmental stage, resulting in a chimeric embryo in which the neural crest cells are histologically detectable. By realizing isochronic (donor and host embryos are at the same developmental stage) and isotopic (the level of the grafted segment corresponds to the level of the excised segment) grafts, the precise map of neural crest derivatives was established, including the migration routes and the levels of origin along the neural tube (see ref. 6 for a review). At the trunk level, neural crest cell migration proceeds mainly in two streams, one dorsoventral near the neural tube within the anterior part of the somite and the other dorsolateral, between ectoderm and somites. The latter pathway is used by melanocyte precursors that migrate in the sub-ectodermal mesenchyme during E3 and E4 of chick development.12,13 Then, other markers have been instrumental to further describe these different migration steps. The monoclonal antibody HNK1/NC114,15 has been widely used as a general marker of neural crest-derived cells in several animal species. It recognizes a glycosylated epitope which is present on most avian crest cells at early stages and later is retained by most crest-derived cells of the peripheral nervous system but not by mesectodermal cells and melanocytes;15 however, HNK1/NC1 reacts with chick (but not quail) melanoblasts at early stages and thus permits following the migration of presumptive quail melanocytes along the dorsolateral pathway in the chick.16 In toto labeling of early chick embryos showed that emigration of melanoblasts from the dorsal neuroepithelium is not segmented. The HNK1positive cells do not enter the lateral space between ectoderm and somites before E3.5. Thereafter they rapidly invade the mediolateral migration pathway located under the ectoderm; they could be found in the middle of the dermomyotome at E4, reaching its lateral edge at E4.5. Then these cells lose HNK1 expression, and their migration cannot be further studied using this antibody. Injection of the lipophilic vital dye DiI into the lumen of the neural tube has been instrumental to marking premigratory neural crest cells and following their migration in vivo. Only the cells present in the neural tube at the time of injection are labeled. Consequently, this technique allowed the successive waves of neural crest cell emigration to be analyzed. Serbedzija et al17 thus performed DiI injections at the trunk level at different developmental stages: Following early injection (at stage 12 of Hamburger and Hamilton) (HH),18 labeled cells were found in all types of neural crest derivatives, including sympathetic ganglia and adrenal medulla; however, in embryos injected at stage 19 HH, only dorsal root ganglia and melanoblasts contained dye-labeled cells; whereas after injection at stage 21 HH, labeled cells were located exclusively along the dorso-lateral pathway. These

Development of Melanocytes from Neural Crest Progenitors

133

experiments thus revealed that as migration proceeds, the distribution of neural crest cells becomes progressively restricted to the more dorsal derivatives, and that late-migrating cells in the trunk only generate melanocytes. Several authors16,17,19,20 noticed that neural crest cells delay their migration before entering the dorso-lateral pathway until this pathway is “opened”. Thus, pigment cell precursors migrate later than other neural crest-derived cells and are still migrating when neural crest-derived cells following the ventral path have already reached their final location in the peripheral nervous system. Even at E5 in avian embryos, Sharma et al21 described neuroepithelial cells capable of migrating and later giving rise to melanocytes. In the mouse, migration of melanoblasts has been first followed indirectly by determining whether isolated fragments of embryonic tissues are capable of giving rise to pigmented cells within a few days. Accordingly, the migration of melanoblasts begins around day 8.5 of gestation and melanoblasts are dispersed throughout the trunk skin ectoderm and adhering mesoderm by day 11.5. After melanoblast colonization is completed (at day 12.5), the precursors go through massive proliferation and start invading the epiderm, in which they will differentiate into melanocytes after birth.5,22,23 More recently DiI was used to trace the first steps of neural crest cell migration in the mouse embryo, in which HNK1 antibody cannot be used.24,25 In the axolotl, neural crest-derived cells seem to first enter the dorso-lateral pathway.26 In Xenopus, neural crest-derived cells migrate also dorsally but pigment cell precursors are found mainly along the ventromedial pathway. In the caudal region, two additional (one dorsal and the other ventral) pathways of presumptive pigment cells have been described using DiI injections.27 In the zebrafish, neurulation proceeds by formation of a neural keel, but the pathways and timing of neural crest cell migration resemble that described in the avian embryo, except that cell migration along the medial pathway is not restricted to the anterior halves of somites and that mesectodermal cells differentiate into cartilage in the trunk, where they give rise to the dorsal fin (reviewed in ref. 28). The zebrafish system allows mutagenesis studies, and thus can provide new information about genes regulating melanocyte development. Two distinct waves of melanoblast migration that depend on different genes have thus been clearly described in the zebrafish. The dorsal epidermal pigment cell precursors migrate precociously and require the spa gene, whereas the dermal pigment cells that migrate later and will form zebra, require ros and leo genes.29 A new field of investigations about the molecular control of pigmentation in the zebrafish is expected from a novel dozen mutants affecting either melanoblast proliferation or melanophore pigmentation pattern.30,31

Potentialities of Neural Crest-derived Cells and Pigment Cell Determination The origin of melanocytes from the pluripotent neural crest cell population raised the question of how differentiation into melanocytes is controlled. Do melanocytes derive from multipotent cells or is the melanocytic lineage segregated early from the other cell types derived from the neural crest? To address these questions, the developmental potential of individual neural crest cells was investigated in culture. The first in vitro cloning analysis32 using a limiting dilution assay revealed 3 types of clones generated by quail trunk neural crest cells: clones that contained only pigment cells or unpigmented cells, and mixed clones with both cell types. Unpigmented cells in mixed colonies were later shown to include several types of neurons.33,34 Therefore pigment cell precursors in the early migratory trunk neural crest comprise both already committed (unipotent) and pluripotent cells. Another method for culturing single neural crest cells was devised in our laboratory, by using single cell plating under microscopic control and culture on a feeder-layer of 3T3 fibroblasts.35 The progeny of migratory neural crest cells isolated from stage 10 HH quail

134

Molecular Basis of Epithelial Appendage Morphogenesis

mes-metencephalon was analyzed with a panel of lineage markers. These studies provided evidence for a large heterogeneity in the proliferation and differentiation potentials of individual crest cells. Most progenitors were found to give rise to two to five different cell types, with various combinations of phenotypes, whereas others generated only glial cells or neurons. In addition, some clones contained all the main phenotypes derived from the cephalic neural crest including cartilage, a mesectodermal derivative, and were therefore considered to derive from a totipotent precursor.35-37 It has to be noted that in these experiments pigment cells were generated exclusively from multipotent (or at least bipotent) cells. Later, a new early marker of avian melanocytes38 allowed unpigmented melanoblasts to be identified in multiphenotypic colonies derived from cephalic and trunk neural crest cells grown on 3T3 fibroblasts.39 Consistent with results of in vivo experiments tracing the fate of single neural crest cells,40 in vitro clonal studies have thus provided evidence that at early migration stages, the neural crest cell population includes pluripotent melanogenic precursors. These results therefore suggested that multipotent cells may be still present at later stages of melanocyte development, and the question was raised of how the final expression of the characteristic phenotypes adopted in each neural crest derivative is controlled. The possibility that crest-derived cells may retain the ability to give rise to pigment cells when they migrate ventrally to form neural derivatives was investigated in vitro. It was shown that in certain conditions pigment cells differentiate from cultured embryonic dorsal root ganglia and peripheral nerves.41-44 It was suggested that melanocytes in these cultures derive from multipotent neural crest cells that exist at early embryonic stages in these locations. Such cells were evidenced by Duff et al45 when they analyzed in vitro the progeny of single cells isolated from quail embryonic peripheral ganglia. Neural crest cells in the epidermis that are normally fated to become melanocytes may also retain multiple developmental potentials. At early stages, they are able to give rise to neurons in vitro46 (Nataf and Le Douarin, unpublished). However, from E6 in quail epiderm, all the crest-derived cells loose their nonmelanogenic developmental options and are determined to the melanocytic phenotype. Artinger and Bronner-Fraser47 distinguished between the first neural crest cells that emigrate from the neural tube in avian embryo at E2 (stage 13 to 18 HH) and those which migrate later (E3-E4) and cannot give rise to adrenergic cells. Therefore, the progressive restriction of neural crest cell fate towards a final differentiation phenotype seems to be a general rule.48,49

Markers of the Melanocytic Lineage Several markers of pigment cell precursors have been produced in the past decade (summarized in Table 8.1). In humans, the melanoblast marker used for developmental studies is the mAb HMB45,50 which detects melanoblasts as early as 40 to 50 days of gestation. Labeling with HMB45 decreases during late gestation and disappears completely after birth.51,52 The glycoprotein labeled by HMB45, p mel17,53 is also recognized by two other antibodies.54 This is one example of the proteins expressed during development and re-expressed during tumor formation. In mouse embryos, melanoblasts are characterized by expression of tyrosinase-related protein 2 (TRP2). TRP2 is the product of the slaty locus55 and catalyzes a step in the melanin synthesis pathway. It is precociously expressed by melanoblasts during embryonic development, as early as 10 days postcoitum (dpc).56,57 Antibodies (ACK2 and ACK4) against the tyrosine kinase receptor c-kit, product of the W locus, were also used as melanoblast markers.58 In avian embryos, three molecules, MEBL-1,59 MelEM38,60 and c-kit,61 were instrumental to follow melanoblast migration, although none of them is entirely specific for the pigment

human

mouse

chick

quail

chick quail mouse

HMB-45

TRP-2

MEBL-1

MelEM

c-kit

mAb: monoclonal antibody

Species

Marker

membrane

cytoplasm

membrane and melanosomes

melanosomes

Pigment Cell Compartment

stage 22 HH

E4

stage 19 HH

12 dpc

40-50 days of gestation

Onset of Labeling

DRG, digestive tract

liver

telencephalon

Other Tissue Labeled

Table 8.1. Characteristics of the different markers for the melanocytic lineage

–

+

+

Pigmented Retina

RNA probe mAb

mAb

mAb

RNA probe

mAb

Type of Reagent

tyrosine-kinase receptor

glutathion S transferase

protein 135 and 115 kDa

tyrosinaserelated protein

glycoprotein

Molecule

Development of Melanocytes from Neural Crest Progenitors 135

136

Molecular Basis of Epithelial Appendage Morphogenesis

cells. The most precocious marker is MEBL1, but it does not label all the pigment cell precursors, since no labeling could be detected in Silky Fowl pigmented internal organs. MelEM is specific for neural crest-derived melanocytes (and does not react with retinal pigmented epithelia) in the quail but stains many other structures in the chick. c-kit is also expressed in dorsal root ganglia.61 When these different markers will be available in the same species, their sequential expression might help in defining different steps of melanoblast differentiation. Furthermore, since some of them are surface markers, cloning analysis of the labeled cells might reveal the state of commitment of these precursors.

Entrance of Melanoblasts in the Epidermis In avian embryos, Le Douarin and Teillet62 constructed quail/chick neuroepithelium chimeras and observed quail neural crest-derived cells entering the dorsal ectoderm of chick hosts as early as E4. Neural crest-derived cells massively invade the epidermis at E5 and this process is completed around E7.13 Hulley et al63 took advantage of the cross between two breeds of chicken whose embryos become pigmented much earlier than other breeds (E5 compared to E7-E9) in order to study the late steps of melanoblast migration. These authors observed that pigmented cells first spread in the dermis at E5. One day later, they are mainly found in the epidermis, and at E8 the nucleus of dermal melanocytes becomes pyknotic. These data therefore indicate that terminal differentiation and migration to the epidermis are not mutually exclusive processes during melanocyte development. In the mouse embryo, recombination experiments between epidermis and dermis were performed to follow epidermis colonization by pigment cell precursors.23 At 10 dpc, melanoblasts are found exclusively in the dermis, and at 11 dpc most of the dermis explants and some of the epidermis explants contained melanoblasts. At 13 and 14 dpc all the epidermal explants were colonized by melanoblasts. Thus, melanoblasts invade the mouse epidermis between 11 and 14 dpc. Recombinations at 14 dpc between the epidermis from albino embryos and the dermis from a pigmented strain did not produce any pigmented hair. This suggests that all the potential sites in ectoderm have been colonized by albino melanoblasts at that stage. Using anti c-kit antibody, Yoshida et al58 showed that melanoblasts go through an active proliferating phase in the dermis before entering the epidermis. Little is known about the mechanisms whereby melanoblasts enter the epidermal layer. Erickson et al16 noticed that neural crest-derived cells invade the epidermis at given places where the basal lamina is disrupted. These authors suggested that those neural crest-derived cells which are the most advanced in the migration process enter the epidermis first. This hypothesis remains to be tested.

Determinants of Migration The existence of pluripotent precursors in the migrating neural crest cell population points out to the importance of the environment in directing their migration and differentiation. Several factors may influence the migration process: extracellular matrix (ECM) and substrates on which neural crest cells migrate, growth and survival factors that control expansion or maintenance of the progenitor pool, and possibly, chemo-attractant factors directing the migration process. Some of these factors have been identified at the molecular level and are described below. Chemo-attractant factors The chemo-attraction hypothesis has been documented.64 When the apical ectodermal ridge is removed from the wing bud or isolated by a membrane, epidermal melano-

Development of Melanocytes from Neural Crest Progenitors

137

blasts are no more concentrated in the distal part of the wing bud but spread all over the wing bud. Other neural crest-derived cells like Schwann cells are not affected by this operation. Thus, epidermal melanoblasts might be attracted by diffusible molecules produced by the apical ectodermal ridge. However, these molecules have not yet been characterized. ECM and adhesion molecules The influence of adhesion molecules on neural crest cell migration has been well documented.65-68 Newgreen and Gooday69 have shown in avian embryos that the beginning of neural crest cell migration coincides with the loss of calcium-dependent adhesion molecules. NCAM, N-cadherin and E-cadherin are no longer expressed when neural crest cells leave the neural tube,70 and the emigration of neural crest cells, as well as their adhesion to the substrate, are dependent on members of the transforming growth factor β (TGF-β) superfamily.71 The presence of numerous ECM components along neural crest migration pathways has been reported: fibronectin,72 laminin,73 collagen,74 and tenascin.75 In vitro, fibronectin and to a lesser extent laminin appear very favorable substrates for neural crest cell migration.76 On the contrary, proteoglycans like chondroitin sulfate are instead inhibitors.77,78 These proteoglycans, such as chondroitin 6 sulfate79 or versican,80 are abundant along the dorso-lateral migration pathway before crest cells start migrating, and disappear when melanoblasts begin to invade the lateral mesenchyme between ectoderm and somites. These molecules might therefore be responsible for the delay of neural crest cell migration along the dorso-lateral pathway.67,80 In axolotl, Spieth and Keller81 showed that adhesion molecules are crucial for pigment cell development: in a white axolotl mutant, the defect of melanoblast migration is due to a delay in the production of a substrate favorable for crest cell migration.82 Moreover, pigmentation of the larva displays alternate bands of xantophores and melanophores. Epperlein and Löfberg83 observed that xantophores migrate later than melanophores. This difference may be due to the fact that only xantophores express NCAM and N-cadherin.84 Growth and survival factors The environment encountered by neural crest-derived cells as they migrate is also crucial through the growth factors allowing survival and expansion of precursor cells. We will rapidly consider here some of these factors whose effect on pigmentation has been reported. Basic fibroblast growth factor (bFGF) was the first melanocyte growth factor identified in pure cultures of human melanocytes.85 In vivo, it is produced by keratinocytes and not by melanocytes, and therefore acts in a paracrine way to regulate pigment cell proliferation.86 In vivo, bFGF is localized in E4 chick embryos in the mesenchyme dorsal to the neural tube where neural crest cells migrate.87 Stocker et al44 showed that bFGF promotes the development of pigment cells in dorsal root ganglia cultures, and reported that TGF-β antagonizes bFGF action. The inhibitory properties of TGF-β on pigment cell differentiation have also been described in quail neural crest cultures.88 Another inhibitory factor, called melanization inhibiting factor (MIF) has been identified in the ventral skin of amphibians.89 This factor may be the amphibian equivalent of the mammalian Agouti Signal Protein (for references, see ref. 90). Platelet-derived growth factor (PDGF) and its receptor PDGFRα were thought to play a role in color patterning, since the Patch mouse mutant harboring a PDGFRα deletion is depigmented.91 However, pigment deficiency is mainly an indirect effect, because the mutation affects primarily the dermis in which melanoblasts migrate and proliferate.92,93 Two other factors of pigment cell development have been identified at the molecular level from genetic studies of coat color in the mouse: the Steel factor, which is the ligand of

138

Molecular Basis of Epithelial Appendage Morphogenesis

the tyrosine kinase c-kit receptor and endothelin 3. They are two examples of genes whose function in pigment cell patterning has been studied in depth. Their discovery and function will be detailed in the next part of this chapter.

From Genes to Mechanisms Regulating Pigmentation A powerful tool for characterizing the molecules that play a role in the onset and maintenance of pigmentation is the analysis of coat color mutations in the mouse, especially those that interfere with melanocyte differentiation during embryonic development. In mice, there are mutations affecting the embryonic development of integumental melanocytes, which typically have no effect on the eyes, but cause a local or global loss of pigmentation from the skin. These loci affect the cells early in their developmental history and interfere with their ability to migrate and colonize the skin. These strains are usually called spotting mutants, in which the degree of pigmentation varies from white spots to complete white coat color (and black eye), the latter being considered as essentially one big spot.94 In this part of the review we would like to describe a decade of research concerning four distinct spotting strains, which led to the identification of two receptor-ligands that were proved to affect melanocytes during two different stages of their development. One receptor-ligand system is c-kit-steel and the other is endothelin receptor B-endothelin 3. Finally we will discuss the possibility that each of these molecular systems corresponds to a different theoretical mode of action long ago proposed to explain spotting.

The C-kit and Steel Molecular System The W and Sl genes encode respectively the c-kit tyrosine kinase receptor and its ligand In the spotting category, two loci with pleiotropic effects on the development of melanocytes, hematopoietic and germ cell lineages have attracted the interest of embryologists for decades. These are: Dominant White spotting (W), identified by Little,95 and Steel (Sl), first described by Sarvella and Russell96 (for reviews see refs. 97, 98). Since then 27 independent alleles have been recognized for the W locus (chromosome 5) and at least 15 different alleles are known for the Sl locus (chromosome 10) (for review, ref. 99). By definition, pigmentation is affected in all of the animals heterozygous for any allele at the W locus, although the extent of white spotting can vary. The characteristic spotting pattern of W heterozygotes is scattered, with no discrete boundaries between pigmented and nonpigmented regions.100 Sl heterozygotes have a slight dilution of the coat color (Steel received its name from this fact) as well as spotting, white blaze on the forehead, white belt spot and unpigmented feet and tail tip.96 No morphological characteristic can be used to definitely distinguish an adult, severely affected but viable, homozygous W/Wv mouse from an adult Sl/Sld mouse since both are black-eyed and completely white. The pigmentation deficiency of W and Sl is caused by the absence of melanocytes in the hair follicles of the coat since the white skin of Wv/Wv and Sl/Sl mice lacks melanocytes, as do the white spots of W and Sl heterozygotes. The eyes are black due to pigmentation in the retina alone; the choroid layer is not pigmented in homozygous, and is “spotted” in heterozygous, mice.101,102 In spite of the phenotypic similarity between W and Sl, the sites and mechanisms of gene action are different. This difference was discovered by various transplantations showing that the defect resides in the stem cell progenitors in W and in the microenvironment through which they develop in Sl.103,104 The complementary sites of action of W and Sl

Development of Melanocytes from Neural Crest Progenitors

139

supported the appealing hypothesis that the genes at these loci encode a receptor and its ligand.97 A major advance which led to the identification of the gene encoded by the W locus was the isolation of the v-kit oncogene as the active transforming gene in a new strain of feline sarcoma virus.105 By homology to kit sequences the human c-kit protooncogene was isolated and characterized. The deduced amino acid sequence of c-kit reveals a cytoplasmic tyrosine-kinase region and a transmembrane glycoprotein containing five extracellular immunoglobulin-like domains that are structurally related to the receptors for CSF-1 and PDGF.106,107 Several groups that looked for a candidate for the W structural gene have demonstrated that c-kit is actually the product of W.108,109 In different W alleles the c-kit gene was found to be either disrupted by genomic rearrangements or to carry a missense mutation that replaces one amino acid in the c-kit protein product.110 In the study of the molecular basis of distinct W phenotypes, Reith et al111 have shown that all the examined W alleles resulted from a deficiency in c-kit associated kinase activity. Mutations that confer reduced levels of an apparently normal protein give mild heterozygous phenotypes, whereas mutations that impair the kinase activity of c-kit give more strongly dominant heterozygous phenotypes. This also suggested that coexpression of normal and defective c-kit proteins in the same cell inhibits proper signal transduction by the wild-type protein. One possible explanation for such a dominant negative mode of action is that c-kit receptors transmit signal by forming dimers and when a wild-type receptor and a functionally inactive receptor form a heterodimer, the signal cannot be transduced. Following the characterization of c-kit in mice and humans, several studies have shown that the gene is expressed and conserved in other species such as rat and chick.112,113 The protein encoded by the Sl locus was identified, on the basis of its expected broad range of biological activities, by several groups114-118 as a transmembrane protein that is processed by proteolytic cleavage in order to release a secreted molecule called here stem cell factor (SCF) or Sl factor. The murine gene was assigned to mouse chromosome 10, the chromosome on which the Sl locus is located,119,120 and SCF sequences were found to be deleted in mice with severe Sl alleles.120 Finally, the hypothesis that Sl and W loci encode a ligand and its receptor97 was challenged. The evidence demonstrating that SCF is the ligand of c-kit came from functional and binding assays.115,116,119 Soluble SCF production is regulated by alternative splicing and is not sufficient for normal development The nucleotide sequence of the cloned SCF cDNA encoded a predicted transmembrane polypeptide.116-118 The SCF protein itself has been identified in two forms. One is a soluble growth factor consisting of the first 164 or 165 aa of the extracellular domain encoded by the cDNA, and the other is a cell surface molecule.114,115,121,122 There are several mechanisms by which a membrane-bound protein could give rise to a soluble protein: 1. the action of a specific or nonspecific protease to cleave the membrane-bound form; 2. alternative splicing of the mRNA to yield a form of the protein that is a better or less efficient substrate for the putative protease; 3. production of a protein that is no longer anchored in the membrane. All these mechanisms were found to be involved in the production of soluble SCF.123-125 However, mice that lack only the transmembrane form of SCF are severely affected by the mutation.123,126 Why the cell surface form of SCF is so critical is unknown, but it was speculated that it might permit a set of direct cell-cell interactions which have a degree of

140

Molecular Basis of Epithelial Appendage Morphogenesis

spatial organization that could not easily be achieved with freely diffused factors. These interactions might be important in processes of normal development, including cellular migration and homing. c-kit and SCF show complementary expression patterns The complementary mode of action of W and Sl loci in the development of the melanocyte stem cell population predicts a complementary expression pattern of their gene products (c-kit and Sl mRNAs, respectively) in this system. More precisely, the cell-autonomous nature of the W mutation and the non cell-autonomous nature of the Sl mutation suggest that the affected cells express c-kit while their microenvironment expresses Sl. In situ expression pattern studies confirmed these predictions. They showed that c-kit and Sl are also expressed in other systems which do not seem to be affected by the mutations and provided additional information, over the mutant phenotype analysis, regarding the developmental stages during which these gene products are needed. In the mouse, the first few cells expressing c-kit mRNA are detected dorsal to the somites within the migratory pathway of neural crest cells at E10, approximately 24-36 h after crest cells emerge from the neural tube. c-kit mRNA positive cells are not detected in the ventral pathway at this time. As development proceeds, c-kit positive cells appear in more and more ventral locations of the dermis and, starting from E13.5, some are already observed in the epidermis. Numerous labeled cells were found in a similar distribution in E15.5 and E17.5. At these stages, however, presumptive melanoblasts could not be distinguished from mast cells, known to express c-kit and to be located in the dermis. At birth, these labeled cells were frequent in the dermis, epidermis and hair follicles.127 The pattern of Sl expression in the developing murine skin is also consistent with a function of this ligand during migration of melanoblasts and their differentiation after cell migration is completed. Sl expression is first detected in the somitic compartment that is the precursor of dermis at a time preceding the influx of pigment cells. At E13, when melanoblast colonization is recently completed in the forelimb, high levels of Sl transcripts are found in mesenchymal cells underlying dermis. The same pattern is observed at E15.5 when melanoblasts are in the process of proliferation and differentiation.128 A role for SCF in melanocyte differentiation was further suggested by the finding that, in transgenic mice, Sl regulatory sequences conferred expression of a LacZ reporter in the dermal papillae of hair follicles.129 In the avian system we could define the exact stage by which neural crest melanocyte precursors start expressing c-kit by using in situ hybridization in combination with the quailchick chimera system.61 After grafting a quail neural primordium into a chick host, neural crest cells could be followed during the entire migration process. We thus found that neural crest cells migrating along the dorsolateral pathway express c-kit as early as E4, that is about two days after the first neural crest cells have left the neural tube. Moreover, the SCF mRNA is expressed from E4 onwards in the epidermis but not in the dermis at any developmental stage. At later stages c-kit positive cells are located at the basal region of the feather buds, whereas high levels of Sl transcripts become restricted to the apical region of the feather buds. This location correlated with the area where melanocytes began to be pigmented at E10.61 At later stages, c-kit and Steel display a complementary expression pattern in the feather barb, suggesting that this system might also play a role in the relationship between melanocytes and keratinocytes (Fig. 8.1). As a whole, these results show that during development c-kit positive cells are present in SCF-producing areas at the time when melanoblasts colonize the embryonic skin. However expression pattern studies do not allow one to determine clearly whether the c-kit/

Development of Melanocytes from Neural Crest Progenitors

141

Fig. 8.1. c-kit and Steel complementary expression in feather filament. In situ hybridization with radioactive riboprobes on transverse sections of E13 chick feather filaments. (A,C) c-kit is expressed in the melanocyte cell bodies which are aligned at the basis of the barb ridges; (B,D) Steel is expressed in the keratinocytes located at the periphery of the barb ridge, which receive the first melanosomes. Thus, the c-kit/Steel system in avians may also play a role in the relationship between melanocytes and keratinocytes. Scale bars represent 26 µm. m: melanocyte, b: barbule cell.

SCF system controls the migration, proliferation and/or differentiation of pigment cell precursors. SCF is required after initial migration and differentiation of melanocyte precursors and before they acquire melanin The first in vivo study investigating the role of c-kit during melanocyte development involved the administration of an anti-c-kit blocking antibody to pregnant mice at different stages of pregnancy.130 At E10.5 and E11.5 the injection of the antibody had basically no effect on coat color of the offspring, and at E12.5 the embryos died in utero due to anemia and thus the effects on pigmentation could not be evaluated. A total depigmentation resulted from injection on E13.5, when melanocyte precursors colonize the skin. During this time, melanoblasts, visualized by anti-c-kit staining, go through a massive proliferation phase in the dermis just before invading the epidermis around E15. From injection on E15.5 no

142

Molecular Basis of Epithelial Appendage Morphogenesis

effect was observed. It was therefore concluded that c-kit may have an essential role for migration and survival/proliferation of the melanoblasts in the dermis until they enter the epidermis. Administration of the antibody during postnatal life revealed that c-kit is also required for melanocyte activation in the hair cycle that occurs throughout life.130 Still the question remained, and could not be answered by these experiments, whether c-kit and SCF are required during the initial stages of melanocyte development. The first indication of the early stages in which the Sl and c-kit gene products are essential for melanocyte precursors came from a comparative study between Sld/Sld and normal mice.56 In this work the TRP-2 probe was used for in situ hybridization to identify melanoblasts in the developing inner ear. TRP-2 was found to detect migratory melanocytes and their precursors as early as E10. In Sld/Sld mice melanoblasts were identified as TRP-2 expressing cells at E11, but there were at least half as many as in normal mice and less advanced in their migration pathway. From E12 onwards, very few melanoblasts were found in the mutants, and none after birth. It seemed, therefore, that the c-kit/Sl system is not needed for the migration and initial differentiation of presumptive melanocytes since melanocyte precursors express TRP-2 before the stage at which they are affected by the mutation. The membrane-bound SCF which is missing in Sld/Sld mice is necessary for the survival/differentiation and targeting of melanoblasts, but not for their early development. The period during which melanocytes require SCF was further investigated using in vitro cultures of murine neural crest cells. The survival of neural crest-derived melanocytes requires SCF for a critical period which begins only after the second day of cell dispersal in vitro and lasts for about 4 days ending at the time that pigment cells differentiate.131 It was also demonstrated that although differentiation does not seem to require SCF, this process may be enhanced by extended exposure to high levels of SCF in vitro.131 These results are in agreement with those previously showing that SCF is required for the maintenance of murine melanocytes.132 In order to define the function of avian SCF we studied the effects of chicken recombinant SCF on the development of melanocytes from quail neural crest cells in vitro. We showed that cultured neural crest cells express the SCF receptor (c-kit mRNA), and that SCF promotes the survival and proliferation of neural crest cells and stimulates the differentiation rate of melanocytic precursors.133 Taken with the expression pattern, we concluded, similarly to studies carried out in mice, that SCF and c-kit are required for the survival and differentiation of melanocyte precursors in the embryonic skin after neural crest cell migration has started.

The Endothelin Molecular System Piebald lethal (sl) and Lethal spotting (ls), two spotting mutations in the mouse Piebald (s locus, chromosome 14) is a recessive mutation known in the mouse for a long time. Homozygotes have dark eyes and show irregular white spotting of the coat, especially on the belly, sides and back. The white areas of the coat completely lack melanocytes. There is a reduction in the number of melanocytes in the choroid layer of the eye. In addition, homozygotes may develop megacolon, which is always associated with lack of ganglion cells in the distal portion of the gut. Mice with a more severe mutation at the piebald locus, piebald lethal (sl), have dark eyes and an almost completely white coat with pigmented hair restricted to small areas on the head and base of the tail. All sl/sl homozygotes develop megacolon with lack of enteric ganglion cells in the posterior end of the colon. They usually die at about two weeks of age, but some live a year or more and may breed.99

Development of Melanocytes from Neural Crest Progenitors

143

Pigment cells and enteric ganglion cells of the colon are both derived from the neural crest, suggesting this structure as the site of action of the s locus. This interpretation was found favorable by experiments investigating the pigmentation and colon innervation deficiencies in these mice. By explantation of embryonic tissues, consisting in the association of neural tubes and skins from wild-type and s/s mice, Mayer22 concluded that the primary cause of spotting in piebald lies in the neural crest. Another recessive mutation that resembles piebald mice is lethal spotting (ls locus on chromosome 2). Lethal spotting mice usually die in the third week of life; however some survive and are fertile. Homozygotes have considerable white spotting and megacolon. As in piebald, the most distal portion of the bowel is aganglionic in ls/ls mice. The absence of the intrinsic reflexes of the enteric nervous system in the terminal gut causes the normally innervated bowel proximal to the aganglionic region to dilate, forming a megacolon. The colons of these mice resemble those of human patients with Hirschsprung’s disease, in which aganglionosis also occurs (see ref. 134 for a review). The deficiency in colon innervation and pigmentation was also studied in ls/ls mice. Mayer and Maltby135 concluded that the ls mutation exerts its effect on pigmentation by reducing the number of melanoblasts. Since melanoblasts and intrinsic ganglion cells are both neural crest derivatives, it is possible that ls acts by causing defective migration or function of these cells or by reducing their number. However, several groups have obtained indirect evidence that in the lower bowel, the ls gene acts by causing an intrinsic abnormality of the terminal 2 mm of the gut.136-138 The s and ls loci encode a receptor and its ligand The breakthrough in the study of the developmental abnormalities caused by sl and ls has been the identification of the genetic defects responsible for these mutations, and the discovery that a related defect is also present in a subset of patients with Hirschsprung’s disease.139-141 These abnormalities are in genes encoding either the endothelin B receptor or its ligand, endothelin 3. The endothelins 1, 2 and 3 (EDN1, EDN2 and EDN3) are a family of 21 aa peptides that activate one or both of the two heptahelical, G-protein-coupled endothelin receptors, A and B (EDNRA and EDNRB). EDNRA exhibits different affinities for endothelin peptides with a potency rank order of EDN1>EDN2>>EDN3.142 EDNRB accepts all three peptides equally,143 as shown in Figure 8.2. Endothelins are each produced from large polypeptide precursors which are first cleaved by furine protease(s) to yield biologically inactive intermediates called big endothelins. Big endothelins, which contain 38 to 41 aa, are further cleaved at a tryptophan residue (Trp-21 site) by endothelin-converting enzymes (ECE).144 This second cleavage produces the active, 21-residue mature forms of endothelins. In ls/ls mice, the gene that encodes EDN3 is mutated: an arginine replaces Trp-21 at the carboxyl terminus of big endothelin 3. Because of this defect, big endothelin 3 cannot be enzymatically converted to the mature active form of EDN3. Moreover, a disruption of the EDN3 gene by homologous recombination leads to an identical ls/ls phenotype of coat color spotting and aganglionic megacolon.141 In sl/sl mice and some humans with Hirschsprung’s disease, missense mutations occur spontaneously in the gene encoding EDNRB.139,140 Moreover, spotting and aganglionic megacolon, identical to that seen in sl/sl mice, develop in animals with homozygous null mutations in EDNRB.140 These studies thus make absolutely clear that both the ligand EDN3 and its receptor EDNRB are necessary for the development of neural crest-derived melanocytes and enteric nerve cells in the terminal bowel. Moreover, the strikingly similar phenotype of mice deficient in either EDN3 or EDNRB suggests that the absence of EDN3 is sufficient to reproduce

144

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 8.2. Schematic representation of the endothelins and their interactions with endothelin receptors. EDNRA binds preferentially EDN1, whereas EDNRB binds the three endothelins with similar affinities.

the EDNRB null phenotype and that EDN1 and EDN2 do not play a major role in the development of these two cell lineages. Further experiments are, however, needed to determine how and when the interaction of EDN3 and EDNRB affects the development of the melanocytic and enteric neuronal lineages from the neural crest. EDN3: A potent mitogen for melanocyte precursors We used an in vitro assay system to study the role of EDN3 during the development of melanogenic cells from the neural crest cell population.145 Addition of EDN3 to quail neural crest cultures strikingly increases the number of melanocytes, which moreover aggregate to form a reproducible pigmentation pattern. Quantitative analysis in different culture conditions revealed that EDN3 significantly promotes neural crest cell proliferation before the appearance of an early marker of melanogenic differentiation. These observations suggest that in vivo EDN3 is essential for an early increase in neural crest cell population to ensure a sufficient supply of melanoblasts and eventually melanocytes in the developing skin.145 The expression pattern of the avian EDN3 receptor (EDNRB) was studied in our laboratory. Neural crest cells start expressing EDNRB at the premigratory stage. Avian EDNRB is thus detected from E2 when crest cells become individualized in the dorsal neural primordium, which is about two days before the onset of c-kit expression. EDNRB transcripts are still present in migratory neural crest cells and later in nerves and ganglia of the peripheral and enteric nervous systems. By contrast, avian neural crest cells migrating in the subectodermal path and, at later stages, melanocytes, fail to express EDNRB.146 However, quail epidermal pigment cells in culture are able to respond to EDN3 stimulation (Dupin, in preparation). Based on this observation, we have recently cloned a novel endothelin receptor subtype expressed in avian pigment cells (Lecoin et al147). The two different receptors might, therefore, account for the two phases of EDN3 response by quail neural crest cells in vitro, i.e., the initial increase in cell proliferation and subsequently the stimulation of melanogenesis. Reid et al148 have described similar effects in vitro of EDN3 on mouse neural crest-derived cells; but in the mouse, EDNRB is expressed by melanocytes (Yanagisawa et al, in preparation).

Development of Melanocytes from Neural Crest Progenitors

145

Mechanisms Leading to Spotting Patterns: Respective Roles of c-kit/Sl and EDNRB/EDN3 In the sixties, mouse geneticists have proposed different models to account for coat color pattern formation. Precise molecular data about the growth factors controlling pigment cell development are now available. We will discuss in this last part how these molecular data might explain the differences in the spotting pattern. As shown in Figure 8.3, the difference is that in c-kit or SCF mutant mice the pigmented areas are scattered or diluted, while the spotted areas in EDNRB or EDN3 deficient mice usually have defined margins. To explain spotting phenotypes, Mintz in 1967149 proposed that the whole coat pigmentation derives from 34 melanoblasts, according to the maximum number of territories that she could observe in aggregation chimeras. Each melanoblast migrates and proliferates along the dorso-ventral axis to populate a stripe-like territory. Spots occur when the descendants of certain clones die, since their progenitors were “preprogrammed” to die.148 Since their death happens relatively late, other clones cannot expand into their territory and an unpigmented region occurs. Schaible examined spotting patterns in different mutants and observed 14 different spotting territories in the coat.149 He therefore suggested that there are 14 melanoblasts, each migrating to the center of a territory and expanding. Spotting occurs when the early melanoblasts die or cannot proliferate. Other clones cannot expand to cover the depigmented area since this process is limited in time. Clones can expand before the tissue environment differentiates to the point that it restricts pigment cell migration.150 More recently Mayer150 suggested that there are natural differences in the distribution of melanogenesis-promoting substances in distinct tissue environments. Thus in areas that are less favorable to melanogenesis, deficient melanoblasts will not survive, resulting in spotting.151 As detailed above, in vitro studies have shown that EDN3 is a potent mitogen for melanocyte progenitors in mouse and avians.145,148 Assuming that EDN3 in vivo also leads to a large expansion of neural crest cells that will eventually differentiate into melanocytes, it is possible that very few precursors bearing the potentiality to give rise to melanocytes exist in the early neural crest cell population as suggested by Mintz and Schaible.149,150,152 The time at which there would be these few precursors probably corresponds to the stage before crest cells need the action of the EDNRB-EDN3 receptor-ligand system. Our data suggest that in avians, between the stages at which melanoblasts require EDNRB and c-kit function, a large expansion in population size occurs. This conclusion agrees with experiments showing that murine EDNRB function is required before E10.5. At that stage, very few melanoblasts are detected in wild-type mice but melanoblasts do not appear in the affected skin areas of sl mice.57 By the time that c-kit is first needed in the mouse (E11.5) the dermis is already seeded with numerous melanoblasts.153 The availability of melanoblast markers (c-kit and TRP-2) allowed Yoshida et al58 to follow directly in whole-mount preparation the distribution of melanoblasts during development of control mouse embryos, ls embryos lacking EDN3, and mouse embryos that had received anti-c-kit blocking antibody at different developing stages. This very elegant study fully confirmed that, when melanoblast development is affected very early (before 12.5 dpc), only some melanoblasts in the cranial and caudal regions that are more advanced in their developmental process escape this impairment to later generate large homogeneous pigmented areas in the head and caudal regions (Fig. 8.3B). By contrast, if melanoblast development is affected at 13.5 dpc when melanoblasts proliferate synchronously before entering the epidermis, the result is an entirely white coat color. Day 14.5 p.c. is critical for colonization of the developing hair follicles, and impairment of melanoblasts at this stage results in a more scattered pattern (Fig. 8.3A).

146

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 8.3. Drawings showing mice with mutation involving c-kit receptor and EDNRB deficiency. Characteristic differences in spotting patterns between (A) a mouse carrying a mutation in the c-kit receptor, with intermingled white and black hair and (B) a mouse with an EDNRB deficiency with sharp margins between white and black regions.

Taken together, spotting in EDNRB or EDN3 mutant mice might occur according to the process provided by Schaible, since the precursors are probably affected before they expand. In contrast, in mice deficient for c-kit or SCF the mechanisms that lead to spotting more resemble those described by Mintz where melanoblasts that are destined to die (i.e., carry a mutation) first expand. These differences might result in the differences in the nature of spots. In EDNRB or EDN3 mutant mice the spots have sharp margins since pigment cell precursors were eliminated early, resulting in the absence of all their descendants in a certain area of the skin. In contrast, in c-kit or SCF deficient mice melanoblasts are affected relatively late. At this stage many melanoblasts are found in the dermis. Partial elimination of melanoblasts would thus result in small distances among pigmented and unpigmented hair. The spotting pattern in mice bearing mutations in both receptor-ligand systems also shares a similar character: in both cases it is the trunk region that is usually depigmented while the head and hindlimb regions often retain pigmentation. Interestingly, it was recently demonstrated that melanoblasts show an uneven distribution along the murine neural axis. In early developmental stages, many more melanocyte precursors are found in the head and tail as compared to the trunk, consistent with those areas that are often pigmented in spotted mice.57,153 The different steps of melanoblast development and the result in terms of color pattern have therefore been fully explored in these two mouse mutants. However, several other spotting mutants whose molecular defects have not yet been identified exist.94 Thus future stud-

Development of Melanocytes from Neural Crest Progenitors

147

ies may supplement our knowledge about pigment cell growth factors. Two other pigmentation loci have been identified: the extension e locus which encodes MSH (melanocyte stimulating hormone) receptor154 and the Agouti A locus155 encoding a paracrine factor which antagonizes MSH (see refs. 156, 157 for a review). These loci regulate the ratio between phaeomelanin (red) and eumelanin (black) and constitute a very nice system which controls the later differentiation steps of pigment cells.

Concluding Remarks Among differentiated cell types, pigment cells have been for long the subject of wide interest to biologists, mostly because of the intriguing variety of pigmentation patterns displayed by adult animals. With the aim of understanding how these patterns are generated during development, many studies were carried out, either by describing in detail the numerous coat color mutants in the laboratory mouse or by exploiting the accessibility of the melanocyte epidermal source to transplantation and grafting experiments. Together with these approaches in mouse genetics and experimental embryology, mostly in avian and amphibian, more recent studies have contributed to understanding the early stages of migration and differentiation of vertebrate pigment cells from their source, the neural crest. Thus the discovery of early markers in different species operationally defined melanoblasts, before their transition to melanocytes expressing tyrosinase and synthesizing pigment granules. Furthermore, the use of powerful culture techniques allowed manipulation of tissues and even single cells in order to investigate how genetic and epigenetic factors control the differentiation of pigment cells from the multipotent neural crest. Recent progress in molecular biology and, particularly, gene targeting techniques provided a molecular explanation for observations made by mouse geneticists several decades ago and suggested mechanisms by which several factors govern the development of pigment cells. So, two molecular systems, the Steel factor and its receptor c-kit, and endothelins and their receptors, have proven to be required for the appearance of melanocytes in the skin. Molecular and functional analysis of the genes in conjunction with cellular approaches in mouse and avian led to a model wherein EDN3/EDNR controls the early proliferation and survival of precursor cells at a stage before Steel/c-kit is required for their survival and maintenance in the developing skin. This sequential view does not exclude the possibility of interactions between EDNRB and c-kit signalings, which remains to be investigated in light of recent findings suggesting a cross-talk between tyrosine-kinase (e.g., c-kit) and G protein-coupled (e.g., EDNRB) receptors in cell lines (see refs. 157, 158 for discussion). Our present knowledge of EDN3 and Steel function does not argue for a role of these factors in directing cells to the pigment phenotype. Therefore a key question still remains as to the molecular mechanism by which neural crest cells become committed to the melanocytic lineage. There again, crucial data could emerge from mouse genetics in conjunction with in vitro cellular techniques. Mi microphtalmia mutant mice with ablated or severely reduced pigmentation harbor mutations in MITF, a transcription factor belonging to the bHLHzipper family of nuclear regulators of differentiation such as MyoD, NeuroD and Mash1.159 The MITF gene has been shown recently to be expressed early in migrating neural crest cells and to be required for their transition to melanoblasts; it therefore may be proposed as a key regulator of cell fate that functions upstream of EDNRB and c-kit to control pigment cell determination.160

Acknowledgments The work in the authors’ laboratory was supported by grants from the Centre National de la Recherche Scientifique, Collège de France, Institut National de la Santé et de la Recherche Médicale, Association pour la Recherche contre le Cancer, and AMGEN Inc. L.L. was

148

Molecular Basis of Epithelial Appendage Morphogenesis

supported in part by the Fondation Bettencourt-Schueller and R.L. was recipient of a fellowship from the Société de Secours des Amis des Sciences. We thank Hélène San Clemente, Francoise Viala and Sophie Gournet for the illustrations, and Marcelle Gendreau for bibliography.

References 1. Du Shane GP. The source of pigment cells in Amphibia. Anat Rec 1934; 60:62-63. 2. Du Shane GP. An experimental study of the origin of pigment cells in Amphibia. J Exp Zool 1935; 72:1-31. 3. Dorris F. Differentiation of pigment cells in tissue cultures of chick neural crest. Proc Soc Exp Biol Med 1936; 34:448-449. 4. Rawles ME. The pigment forming potency of early chick blastoderm. Proc Natl Acad Sci USA 1940a; 26:86-94. 5. Rawles M. The development of melanophores from embryonic mouse tissue grown in the coelom of chick embryos. Proc Natl Acad Sci USA 1940b; 26:673-680. 6. Le Douarin NM. The neural crest. 259p. Cambridge University Press, Cambridge.1982. 7. Watterson RL. On the production of feather color pattern by mesodermal grafts between Barred Plymouth Rock and White Leghorn chick embryos. Anat Rec Suppl 1938; 72:100-101. 8. Eastlick HL. The point of origin of the melanophores in chick embryos as shown by means of limb bud transplants. J Exp Zool 1939; 82:131-159. 9. Willier BH, Rawles ME. The control of feather color pattern by melanophores grafted from one embryo to another of a different breed of fowl. Physiol Zool 1940; 13:177-202. 10. Ris H. An experimental study of the origin of melanophores in birds. Physiol Zool 1941; 14:48-66. 11. Le Douarin N. Particularités du noyau interphasique chez la Caille japonaise (Coturnix coturnix japonica). Utilisation de ces particularités comme “marquage biologique” dans les recherches sur les interactions tissulaires et les migrations cellulaires au cours de l’ontogenèse. Bull Biol Fr Belg 1969; 103:435-452. 12. Teillet MA, Le Douarin NM. La migration des cellules pigmentaires étudiée par la methode des greffes hétérospecifiques de tube nerveux chez l’embryon d’oiseau. C R Acad Sci, Serie III, Paris 1970; 270:3095-3098. 13. Teillet MA. Recherches sur le mode de migration et la différenciation des mélanoblastes cutanés chez l’embryon d’oiseau: Etude expérimentale par la méthode des greffes hétérospécifiques entre embryons de Caille et de Poulet. Ann Embryol Morphog 1971; 1971:95-109. 14. Abo T, Balch CM. A differentiation antigen of human NK and K cells identified by a monoclonal antibody (HNK-1). J Immunol 1981; 121:1024-1029. 15. Vincent M, Duband JL, Thiery JP. A cell surface determinant expressed early on migrating avian neural crest cells. Dev Brain Res 1983; 9:235-238. 16. Erickson CA, Duong TD, Tosney KW. Descriptive and experimental analysis of the dispersion of neural crest cells along the dorsolateral path and their entry into ectoderm in the chick embryo. Dev Biol 1992; 151:251-272. 17. Serbedzija GN, Bronner-Fraser M, Fraser SE. A vital dye analysis of the timing and pathways of avian trunk neural crest cell migration. Development 1989; 106:809-816. 18. Hamburger V, Hamilton HL. A series of normal stages in the development of chick embryo. J Morphol 1951; 88:49-92. 19. Weston JA. A radioautographic analysis of the migration and localization of trunk neural crest in the chick. Dev Biol 1963; 6:279-310. 20. Tosney KW. The early migration of neural crest cells in the trunk region of the avian embryo: an electron microscopic study. Dev Biol 1978; 62:317-333. 21. Sharma K, Korade Z, Frank E. Late-migrating neuroepithelial cells from the spinal cord differentiate into sensory ganglion cells and melanocytes. Neuron 1995; 14:143-152. 22. Mayer TC. The development of piebald spotting in mice. Dev Biol 1965; 11:319-334.

Development of Melanocytes from Neural Crest Progenitors

149

23. Mayer TC. The migratory pathway of neural crest cells into the skin of mouse embryos. Dev Biol 1973; 34:39-46. 24. Serbedzija GN, Fraser SE, Bronner-Fraser M. Pathways of trunk neural crest cell migration in the mouse embryo as revealed by vital dye labelling. Development 1990; 108:605-612. 25. Serbedzija GN, Bronner-Fraser M, Fraser SE. Vital dye analysis of cranial neural crest cell migration in the mouse embryo. Development 1992; 116:297. 26. Löfberg J, Ahlfors K, Föllström C. Neural crest cell migration in relation to extracellular matrix organization in the embryonic axolotl trunk. Dev Biol 1980; 75:148-167. 27. Collazo A, Bronner-Fraser M, Fraser SE. Vital dye labelling of Xenopus laevis trunk neural crest reveals multipotency and novel pathways of migration. Development 1993; 118:363-376. 28. Eisen JS, Weston JA. Development of the neural crest in the zebrafish. Dev Biol 1993; 159:50-59. 29. Johnson SL, Africa D, Walker C et al. Genetic control of adult pigment stripe development in zebrafish. Dev Biol 1995; 167:27-33. 30. Kelsh RN, Brand M, Jiang YJ et al. Zebrafish pigmentation mutations and the processes of neural crest development. Development 1996; 123:369-389. 31. Odenthal J, Rossnagel K, Haffter P et al. Mutation affecting xanthophores pigmentation in the zebrafish, Danio rerio. Development 1996; 123:391-398. 32. Cohen AM, Konigsberg IR. A clonal approach to the problem of neural crest determination. Dev Biol 1975; 46:262-280. 33. Sieber-Blum M, Cohen AM. Clonal analysis of quail neural crest cells: they are pluripotent and differentiate in vitro in the absence of noncrest cells. Dev Biol 1980; 80:96-106. 34. Sieber-Blum M. Commitment of neural crest cells to the sensory neuron lineage. Science 1989; 243:1608-1611. 35. Baroffio A, Dupin E, Le Douarin NM. Clone-forming ability and differentiation potential of migratory neural crest cells. Proc Natl Acad Sci USA 1988; 85:5325-5329. 36. Dupin E, Baroffio A, Dulac C et al. Schwann-cell differentiation in clonal cultures of the neural crest, as evidenced by the anti-Schwann cell myelin protein monoclonal antibody. Proc Natl Acad Sci USA 1990; 87:1119-1123. 37. Baroffio A, Dupin E, Le Douarin NM. Common precursors for neural and mesectodermal derivatives in the cephalic neural crest. Development 1991; 112:301-305. 38. Nataf V, Mercier P, Ziller C et al. Novel markers of melanocyte differentiation in the avian embryo. Exp Cell Res 1993; 207:171-182. 39. Dupin E, Le Douarin NM. Retinoic acid promotes the differentiation of adrenergic cells and melanocytes in quail neural crest cultures. Dev Biol 1995; 168:529-548. 40. Bronner-Fraser M, Fraser SE. Cell lineage analysis reveals multipotency of some avian neural crest cells. Nature 1988; 335:161-164. 41. Nichols DH, Weston JA. Melanogenesis in cultures of peripheral nervous tissue. I. The origin and prospective fate of cells giving rise to melanocytes. Dev Biol 1977; 60:217-225. 42. Nichols DH, Kaplan RA, Weston JA. Melanogenesis in cultures of peripheral nervous tissue. II. Environmental factors determining the fate of pigment-forming cells. Dev Biol 1977; 60:226-237. 43. Ciment G, Glimelius B, Nelson DM et al. Reversal of a developmental restriction in neural crest-derived cells of avian embryos by a phorbol ester drug. Dev Biol 1986; 118:392-398. 44. Stocker KM, Sherman L, Rees S et al. Basic FGF and TGF-beta 1 influence commitment to melanogenesis in neural crest-derived cells of avian embryos. Development 1991; 111:635-645. 45. Duff RS, Langtimm CJ, Richardson MK et al. In vitro clonal analysis of progenitor cell patterns in dorsal root and sympathetic ganglia of the quail embryo. Dev Biol 1991; 147:451-459. 46. Richardson MK, Sieber-Blum M. Pluripotent neural crest cells in the developing skin of the quail embryo. Dev Biol 1993; 157:348-358. 47. Artinger KB, Bronner-Fraser M. Partial restriction in the developmental potential of late emigrating avian neural crest cells. Dev Biol 1992; 149:149-157.

150

Molecular Basis of Epithelial Appendage Morphogenesis

48. Sextier-Sainte-Claire Deville F, Ziller C, Le Douarin NM. Developmental potentialities of cells derived from the truncal neural crest in clonal cultures. Dev Brain Res 1992; 66:1-10. 49. Sextier-Sainte-Claire Deville F, Ziller C, Le Douarin NM. Developmental potentials of enteric neural crest-derived cells in clonal and mass cultures. Dev Biol 1994; 163:141-151. 50. Gown AM, Vogel AM, Hoak D et al. Monoclonal antibodies specific for melanocytic tumors distinguish subpopulations of melanocytes. Am J Pathol 1986; 123:195-203. 51. Holbrook KA, Vogel AM, Underwood RA et al. Melanocytes in human embryonic and fetal skin: a review and new findings. Pigment Cell Res Suppl 1988; 1:6-17. 52. Holbrook KA, Underwood RA, Vogel AM et al. The appearance, density and distribution of melanocytes in human embryonic and fetal skin revealed by the anti-melanoma monoclonal antibody, HMB-45. Anat Embryol 1989; 180:443-455. 53. Kwon BS, Chintamaneni C, Kozak CA et al. A melanocyte-specific gene, Pmel 17, maps near the silver coat color locus on mouse chromosome 10 and is in a syntenic region on human chromosome 12. Proc Natl Acad Sci USA 1991; 88:9228-9232. 54. Adema GJ, de Boer AJ, Van ‘T Hullenaar R et al. Melanocyte lineage-specific antigens recognized by monoclonal antibodies NKI-beteb, HMB-50, and HMB-45 are encoded by a single cDNA. Am J Pathol 1993; 143:1579-1585. 55. Jackson IJ, Chambers DM, Tsukamoto K et al. A second tyrosinase-related protein, TRP-2, maps to and is mutated at the mouse slaty locus. EMBO J 1992; 11:527-535. 56. Steel KP, Davidson DR, Jackson IJ. TRP-2/DT, a new early melanoblast marker, shows that steel growth factor (c-kit ligand) is a survival factor. Development 1992; 115:1111-1119. 57. Pavan WJ, Tilghman SM. Piebald lethal (sl) acts early to disrupt the development of neural crest-derived melanocytes. Proc Natl Acad Sci USA 1994; 91:7159-7163. 58. Yoshida H, Kunisada T, Kusakabe M et al. Distinct stages of melanocyte differentiation revealed by analysis of nonuniform pigmentation patterns. Development 1996; 122: 1207-1214. 59. Kitamura K, Takiguchi-Hayashi K, Sezaki M et al. Avian neural crest cells express a melanogenic trait during early migration from the neural tube: observations with the new monoclonal antibody, MEBL-1. Development 1992; 114:367-378. 60. Nataf V, Mercier P, De Nechaud B et al. Melanoblast/ Melanocyte early marker (MelEM) is a glutathione S-transferase subunit. Exp Cell Res 1995; 218:394. 61. Lecoin L, Lahav R, Martin FH et al. Steel and c-kit in the development of avian melanocytes: A study of normally pigmented birds and of the hyperpigmented mutant silky fowl. Dev Dyn 1995; 203:106-118. 62. Le Douarin NM, Teillet MA. Sur quelques aspects de la migration des cellules neurales chez l’embryon de poulet etudiee par la methode des greffes heterospecifiques de tube nerveux. C R Soc Biol 1970; 164:390-397. 63. Hulley PA, Stander CS, Kidson SH. Terminal migration and early differentiation of melanocytes in embryonic chick skin. Dev Biol 1991; 145:182-194. 64. Wachtler F. On the migration of epidermal melanoblasts in the avian embryonic wing bud. Anat Embryol 1984; 170:307-312. 65. Bilozur ME, Hay ED. Neural crest migration in 3D extracellular matrix utilizes laminin, fibronectin, or collagen. Dev Biol 1988; 125:19-33. 66. Perris R, Von Böxberg Y, Lofberg J. Local embryonic matrices determine region-specific phenotypes in neural crest cells. Science 1988; 241:86-89. 67. Erickson CA, Perris R. The role of cell-cell and cell-matrix interactions in the morphogenesis of the neural crest. Dev Biol 1993; 159:60-74. 68. Perris R. The extracellular matrix in neural crest-cell migration. Trends in Neurosci 1997; 20:23-31. 69. Newgreen DF, Gooday D. Control of the onset of migration of neural crest cells in avian embryos. Role of Ca++-dependent cell adhesions. Cell Tissue Res 1985; 239:329-336. 70. Duband JL, Volberg T, Sabanay I et al. Spatial and temporal distribution of the adherensjunction-associated adhesion molecule A-CAM during avian embryogenesis. Development 1988; 103:325-344.

Development of Melanocytes from Neural Crest Progenitors

151

71. Delannet M, Duband JL. Transforming growth factor-beta control of cell-substratum adhesion during avian neural crest cell emigration in vitro. Development 1992; 116:275-287. 72. Newgreen D, Thiery JP. Fibronectin in early avian embryos: synthesis and distribution along the migration pathways of neural crest cells. Cell Tiss Res 1980; 211:269-291. 73. Rogers SL, Edson KJ, Letourneau PC et al. Distribution of laminin in the developing peripheral nervous system of the chick. Dev Biol 1986; 113:429-435. 74. Duband JL, Thiery JP. Distribution of laminin and collagens during avian neural crest development. Development 1987; 101:461-478. 75. Epperlein HH, Halfter W, Tucker RP. The distribution of fibronectin and tenascin along migratory pathways of the neural crest in the trunk of amphibian embryos. Development 1988; 103:743-756. 76. Rovasio RA, Delouvee A, Yamada KM et al. Neural crest cell migration: requirements for exogenous fibronectin and high cell density. J Cell Biol 1983; 96:462-473. 77. Tan SS, Crossin KL, Hoffman S et al. Asymmetric expression in somites of cytotactin and its proteoglycan ligand is correlated with neural crest cell distribution. Proc Natl Acad Sci USA 1987; 84:7977-7981. 78. Perris R, Krotoski D, Lallier T et al. Spatial and temporal changes in the distribution of proteoglycans during avian neural crest development. Development 1991; 111:583-599. 79. Oakley RA, Lasky CJ, Erickson CA et al. Glycoconjugates mark a transient barrier to neural crest migration in the chicken embryo. Development 1994; 120:103-114. 80. Landolt RM, Vaughan L, Winterhalter KH et al. Versican is selectively expressed in embryonic tissues that act as barriers to neural crest cell migration and axon outgrowth. Development 1995; 121:2303-2312. 81. Spieth J, Keller RE. Neural crest cell behavior in white and dark larvae of Ambystoma mexicanum: differences in cell morphology, arrangement, and extracellular matrix as related to migration. J Exp Zool 1984; 229:91-107. 82. Keller RE, Löfberg J, Spieth J. Neural crest cell behavior in white and dark embryos of Ambystoma mexicanum: epidermal inhibition of pigment cell migration in the white axolotl. Dev Biol 1982; 89:179-195. 83. Epperlein HH, Löfberg J. Xantophores in chromatophores groups of the premigratory neural crest initiate the pigment pattern of the axolotl larva. Roux’s Arch Dev Biol 1984; 193:357-369. 84. Fukusawa T, Obika M. NCAM and N-cadherin are specifically expressed in xantophores, but not in the other types of pigment cells, melanophores, and iridophores. Pigment Cell Res 1995; 8:1-9. 85. Halaban R, Ghosh S, Baird A. bFGF is the putative natural growth factor for human melanocytes. In Vitro Cell Dev Biol 1987; 23:47-52. 86. Halaban R, Langdon R, Birchall N et al. Basic fibroblast growth factor from human keratinocytes is a natural mitogen for melanocytes. J Cell Biol 1988; 107:1611-1619. 87. Kalcheim C, Neufeld G. Expression of basic fibroblast growth factor in the nervous system of early avian embryos. Development 1990; 109:203-215. 88. Rogers SL, Gegick PJ, Alexander SM et al. Transforming growth factor-beta alters differentiation in cultures of avian neural crest-derived cells: effects on cell morphology, proliferation, fibronectin expression, and melanogenesis. Dev Biol 1992; 151:192-203. 89. Fukuzawa T, Ide H. A ventrally localized inhibitor of melanization in Xenopus laevis skin. Dev Biol 1988; 129:25-36. 90. Houillon C, Bagnara JT. Insights into pigmentary phenomena provided by grafting and chimera formation in the axolotl. Pigment Cell Res 1996; 9:281-288. 91. Stephenson DA, Mercola M, Anderson E et al. Platelet-derived growth factor receptor alpha-subunit gene (Pdgfrα) is deleted in the mouse patch (Ph) mutation. Proc Natl Acad Sci USA 1991; 88:6-10. 92. Morrison-Graham K, Schatteman GC, Bork T et al. A PDGF receptor mutation in the mouse (Patch) perturbs the development of a non-neuronal subset of neural crest-derived cells. Development 1992; 115:133-142.

152

Molecular Basis of Epithelial Appendage Morphogenesis

93. Orr-Urtreger A, Bedford MT, Do MS et al. Developmental expression of the alpha receptor for platelet-derived growth factor, which is deleted in the embryonic lethal Patch mutation. Development 1992; 115:289-303. 94. Bennett DC. Genetics, development, and malignancy of melanocytes. In: Jeon KW, Jarvik J, Friedlander M, eds. International Review of Cytology: A Survey of Cell Biology, vol 146. San Diego: Academic Press, 1993:191-260. 95. Little CC. The inheritance of black-eyed white spotting in mice. Am Nat 1915; 49:727-740. 96. Sarvella PA, Russell LB. Steel, a new dominant gene in the house mouse with effects on coat pigment and blood. J Heredity 1956; 47:123-128. 97. Russell, E. Hereditary anemias of the mouse: a review for geneticists. Adv Genet 1979; 20:357-459. 98. Silvers W. 1979. White spotting: piebald, lethal spotting, and Belted. The coat color of Mice: a model for gene action and interaction (W. Silvers), Springer Verlag, New York. 99. Lyon MF, Searle AG. 1990. Genetic variants and strains of the laboratory mouse. Oxford University Press. 100. Geissler EN, McFarland EC, Russell ES. Analysis of pleiotropism at the dominant whitespotting (W) locus of the house mouse: a description of ten new W alleles. Genetics 1981; 97:337-361. 101. Markert CL, Sievers WK. The effects of genotype and cell environment on melanoblast differentiation in the house mouse. Genetics 1956; 41:429-450. 102. Bennett D. Developmental analysis of a mutation with pleiotropic effect in the mouse. J Morphol 1956; 98:199-233. 103. Russell E, Smith LJ, Lawson FA. Implantation of normal blood-forming tissue in radiated genetically anemic hosts. Science 1956; 124:1076-1077. 104. Mayer TC, Green TC. An experimental analysis of the pigment defect caused by mutations at the W and Sl loci in mice. Dev Biol 1968; 18:62-75. 105. Besmer P, Murphy JE, George PC et al. A new acute transforming feline retrovirus and relationship of its oncogene v-kit with the protein kinase gene family. Nature 1986; 320:415-421. 106. Yarden Y, Kuang WJ, Yang-Feng T et al. Human proto-oncogene c-kit: a new cell surface receptor tyrosine kinase for an unidentified ligand. EMBO J 1987; 6:3341-3351. 107. Qiu FH, Ray P, Brown K et al. Primary structure of c-kit: relationship with the CSF-1/ PDGF receptor kinase family-oncogenic activation of v-kit involves deletion of extracellular domain and C terminus. EMBO J 1988; 7:1003-1011. 108. Chabot B, Stephenson DA, Chapman VM et al. The proto-oncogene c-kit encoding a transmembrane tyrosine kinase receptor maps to the mouse W locus. Nature 1988; 335:88-89. 109. Geissler EN, Ryan MA, Housman E. The dominant-white spotting (W) locus of the mouse encodes the c-kit proto-oncogene. Cell 1988; 55:185-192. 110. Tan JC, Nocka K, Ray P et al. The dominant W42 spotting phenotype results from a missense mutation in the c-kit receptor kinase. Science 1990; 247:209-212. 111. Reith AD, Rottapel R, Giddens E et al. W mutant mice with mild or severe developmental defects contain distinct point mutations in the kinase domain of the c-kit transmembrane receptor. Genes Dev 1990; 4:390-400. 112. Tsujimura T, Hirota S, Nomura S et al. Characterization of Ws mutant allele of rats: a 12 base deletion in tyrosine kinase domain of c-kit gene. Blood 1991; 78:1942-1946. 113. Sasaki E, Okamura H, Chikamune T et al. Cloning and expression of the chicken c-kit proto-oncogene. Gene 1993; 128:257-261. 114. Zsebo KM, Williams DA, Geissler EN et al. Stem cell factor is encoded at the Sl locus of the mouse and is the ligand for the c-kit tyrosine kinase receptor. Cell 1990; 63:213-224. 115. Williams DE, Eisenman J, Baird A et al. Identification of a ligand for the c-kit protooncogene. Cell 1990; 63:167-174. 116. Huang E, Nocka K, Beier DR et al. The hematopoietic growth factor KL is encoded by the Sl locus and is the ligand of the c-kit receptor, the gene product of the W locus. Cell 1990; 63:225-233.

Development of Melanocytes from Neural Crest Progenitors

153

117. Martin FH, Suggs SV, Langley KE et al. Primary structure and functional expression of rat and human stem cell factor DNAs. Cell 1990; 63:203-211. 118. Anderson DM, Lyman SD, Baird A et al. Molecular cloning of mast cell growth factor, a hematopoietin that is active in both membrane bound and soluble. Cell 1990; 63:235-243. 119. Zsebo KM, Wypych J, McNiece IK et al. Identification, purification, and biological characterization of hematopoietic stem cell factor from buffalo rat liver-conditioned medium. Cell 1990; 63:195-201. 120. Copeland NG, Gilbert DJ, Cho BC et al. Mast cell growth factor maps near the steel locus on mouse chromosome 10 and is deleted in a number of steel alleles. Cell 1990; 63:175-183. 121. Nocka K, Buck J, Levi E et al. Candidate ligand for the c-kit transmembrane kinase receptor: KL, a fibroblast derived growth factor stimulates mast cells and erythroid progenitors. EMBO J 1990; 9:3287-3294. 122. Flanagan JG, Leder P. The kit ligand: a cell surface molecule altered in steel mutant fibroblasts. Cell 1990; 63:185-194. 123. Flanagan JG, Chan DC, Leder P. Transmembrane form of the kit ligand growth factor is determined by alternative splicing and is missing in the Sld mutant. Cell 1991; 64:1025-135. 124. Huang EJ, Nocka KH, Buck J et al. Differential expression and processing of two cell associated forms of the kit-ligand: KL-1 and KL-2. Mol Biol Cell 1992; 3:349-362. 125. Majumdar MK, Feng L, Medlock E et al. Identification and mutation of primary and secondary proteolytic cleavage sites in murine stem cell factor cDNA yields biologically active, cell-associated protein. J Biol Chem 1994; 269:1237-1242. 126. Brannan CI, Bedell MA, Resnick JL et al. Developmental abnormalities in Steel17H mice result from a splicing defect in the steel factor cytoplasmic tail. Genes Dev 1992; 6:1832-1842. 127. Manova K, Bachvarova RF. Expression of c-kit encoded at the W locus of mice in developing embryonic germ cells and presumptive melanoblasts. Dev Biol 1991; 146:312-324. 128. Keshet E, Lyman SD, Williams DE et al. Embryonic RNA expression patterns of the c-kit receptor and its cognate ligand suggest multiple functional roles in mouse development. EMBO J 1991; 10:2425-2435. 129. Yoshida H, Hayashi S, Shultz LD et al. Neural and skin-specific expression pattern conferred by steel factor regulatory sequence in transgenic mice. Dev Dyn 1996; 207:222-232. 130. Nishikawa S, Kusakabe M, Yoshinaga K et al. In utero manipulation of coat color formation by a monoclonal anti-c-kit antibody: two distinct waves of c-kit-dependency during melanocyte development. EMBO J 1991; 10:2111-2118. 131. Morrison-Graham K, Weston JA. Transient steel factor dependence by neural crest-derived melanocyte precursors. Dev Biol 1993; 159:346-352. 132. Murphy M, Reid K, Williams DE et al. Steel factor is required for maintenance, but not differentiation, of melanocyte precursors in the neural crest. Dev Biol 1992; 153:396-401. 133. Lahav R, Lecoin L, Ziller C et al. Effect of the Steel gene product on melanogenesis in avian neural crest cell cultures. Differentiation 1994; 58:133-139. 134. Gershon MD. Neural crest development—Do developing enteric neurons need endothelins? Curr Biol 1995; 5:601-604. 135. Mayer TC, Maltby EL. An experimental investigation of pattern development in lethal spotting and belted mouse embryos. Dev Biol 1964; 9:269-286. 136. Rothman TP, Gershon MD. Regionally defective colonization of the terminal bowel by the precursors of enteric neurons in lethal spotted mutant mice. Neuroscience 1984; 12:1293-1311. 137. Kapur RP, Yost C, Palmiter RD. Aggregation chimeras demonstrate that the primary defect responsible for aganglionic megacolon in lethal-spotted mice is not neuroblast autonomous. Development 1993; 117:993-999. 138. Rothman TP, Goldowitz D, Gershon MD. Inhibition of migration of neural crest-derived cells by the abnormal mesenchyme of the presumptive aganglionic bowel of ls/ls mice: analysis with aggregation and interspecies chimeras. Dev Biol 1993; 159:559-573. 139. Puffenberger EG, Hosoda K, Washington SS et al. A missense mutation of the endothelinB receptor gene in multigenic Hirschsprung’s disease. Cell 1994; 79:1257-1266.

154

Molecular Basis of Epithelial Appendage Morphogenesis

140. Hosoda K, Hammer RE, Richardson JA et al. Targeted and natural (piebald-lethal) mutations of endothelin-B receptor gene produce megacolon associated with spotted coat color in mice. Cell 1994; 79:1267-1276. 141. Greenstein-Baynash A, Hosoda K, Giaid A et al. Interaction of endothelin-3 with endothelinB receptor is essential for development of neural crest-derived melanocytes and enteric neurons: Missense mutation of endothelin-3 gene in lethal spotting mice. Cell 1994; 79:277-1285. 142. Yanagisawa M. The endothelin system A new target for therapeutic intervention. Circulation 1994; 89:320-1322. 143. Sakurai T, Yanagisawa M, Takuwa Y et al. Cloning of a cDNA encoding a non-isopeptideselective subtype of the endothelin receptor. Nature 1990; 348:32-735. 144. Xu D, Emoto N, Giaid A et al. ECE-1: A membrane-bound metalloprotease that catalyzes the proteolytic activation of big endothelin-1. Cell 1994; 78:73-485. 145. Lahav R, Ziller C, Dupin E et al. Endothelin 3 promotes neural crest cell proliferation and mediates a vast increase in melanocyte number in culture. Proc Natl Acad Sci USA 1996; 93:3892-3897. 146. Nataf V, Lecoin L, Lahav R et al. Endothelin-B receptor is expressed by neural crest cells in the avian embryo. Proc Natl Acad Sci 1996; 93:9645-9650. 147. Lecoin L, Sakurai T, Ngo MT et al. Cloning and characterization of a novel endothelin receptor subtype in the avian class. Proc Natl Acad Sci USA 1998; 95:3024-3029. 148. Reid K, Turnley AM, Maxwell GD et al. Multiple roles of endothelin in melanocyte development: Regulation of progenitor number and stimulation of differentiation. Development 1996; 122:3911-3919. 149. Mintz B. Gene control of mammalian pigmentary differentiation. I. Clonal origin of melanocytes. Proc Natl Acad Sci USA 1967; 58:344-351. 150. Schaible RH. Clonal distribution of melanocytes in piebald-spotted and variegated mice. J Exp Zool 1969; 172:181-200. 151. Mayer TC. Enhancement of melanocyte development from piebald neural crest by a favorable tissue environment. Dev Biol 1977; 56:255-262. 152. Mintz B. A comparison of pigment cell development in albino, steel and dominant-spotting mutant mouse embryos. Dev Biol 1967; 23:297-309. 153. Wehrle-Haller B, Weston JA. Soluble and cell-bound forms of steel factor activity play distinct roles in melanocyte precursor dispersal and survival on the lateral neural crest migration pathway. Development 1995; 121:731-742. 154. Robbins LS, Nadeau JH, Johnson KR et al. Pigmentation phenotypes of variant extension locus alleles result from point mutations that alter MSH receptor function. Cell 1993; 72:827-834. 155. Miller MW, Duhl DM, Vrieling H et al. Cloning of the mouse agouti gene predicts a secreted protein ubiquitously expressed in mice carrying the lethal yellow mutation. Genes Dev 1993; 7:454-467. 156. Millar SE, Miller MW, Stevens ME et al. Expression and transgenic studies of the mouse agouti gene provide insight into the mechanisms by which mammalian coat color patterns are generated. Development 1995; 121:3223-3232. 157. Barsh GS. The genetics of pigmentation: From fancy genes to complex traits. Trends in Genetics 1996; 12:299-305. 158. Daub H, Weiss FU, Wallasch C et al. Role of transactivation of the EGF receptor in signalling by G-protein-coupled receptors. Nature 1996; 379;557-560. 159. Moore KJ. Insight into the microphtalmia gene. Trends in Genetics 1995; 11:442-448. 160. Opdecamp K, Nakayama A, Nguyen MT et al. Melanocyte development in vivo and in neural crest cell cultures: Crucial dependence on the Mitf basic-helix-loop-helix-zipper transcription factor. Development 1997; 124:2377-2386.

Development of Melanocytes from Neural Crest Progenitors

Part III

Morphogenesis of Epithelial Appendages Within the Body

155

CHAPTER 9

The Teeth as Models for Studies on the Molecular Basis of the Development and Evolution of Organs Irma Thesleff and Johanna Pispa

Introduction

T

eeth develop from stomodeal or pharyngeal epithelium and the underlying neural crestderived mesenchymal cells in a way very similar to skin derivatives such as hair, feathers and scales (Fig. 9.1). The first morphological sign of tooth development is a thickening of the oral epithelium, which subsequently buds into the underlying mesenchyme. The mesenchymal cells condense around the bud, and during the following cap and bell stages the epithelium undergoes folding morphogenesis, resulting in the establishment of the tooth crown form. The mesenchyme, which becomes surrounded by the dental epithelium, forms the dental papilla, giving rise to tooth pulp and the odontoblasts. The more peripheral cells of the condensed dental mesenchyme extend around the epithelial dental component, the enamel organ, forming the dental follicle. The follicle subsequently gives rise to the cementoblasts depositing dental cementum, as well as to the fibrous periodontal membrane connecting the roots of the teeth to the alveolar bone. It should be noted that teeth always develop separately from bones, although in fish, amphibians and some reptiles they become united to bone during development. In many vertebrates, notably in mammals, the periodontal membrane separates the teeth from the bones, and the teeth have a central role in the regulation of the growth of the alveolar bone. The terminal differentiation of the dentine-forming odontoblasts and the enamel-forming ameloblasts takes place during the bell stage at the interface of the epithelium and mesenchyme. The preodontoblast cells align under the epithelial basement membrane, polarize and start to secrete the collagenous dentin matrix, which subsequently mineralizes into a bone-like hard tissue. The preameloblast cells opposite the odontoblasts become columnar, their nuclei polarize and they start the deposition of the enamel matrix. This matrix directs the mineralization of the enamel into the hardest tissue in the body, and during this process the enamel proteins are almost entirely degraded. After crown morphogenesis, the roots of the teeth develop and subsequently the teeth erupt into the oral cavity (Fig. 9.1, for more details see ref. 1). It is apparent that the morphological features of early tooth development, as well as the molecular mechanisms regulating tooth morphogenesis, bear more similarities than differences to the development of other epithelial appendages2 (see chapter 5). In fact, no Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

Molecular Basis of Epithelial Appendage Morphogenesis

158

Fig. 9.1. Tooth development. Tooth formation is initiated by the thickening of the oral epithelium, which is followed by the epithelium budding into the mesenchyme, with mesenchymal cells condensing around the tooth bud. During cap stage the epithelium folds and surrounds the condensed mesenchymal cells which form the dental papilla. The epithelium continues to fold at the bell stage, resulting in the formation of the final tooth shape. Differentiated epithelial (ameloblast) and mesenchymal (odontoblast) cells start to secrete enamel and dentin, respectively. Roots develop after the crown is formed, and finally the tooth erupts into the oral cavity.

developmental mechanisms or regulatory molecules have so far been shown to be really unique for tooth development. A particular feature of tooth development, which is shared by hair development, is that morphogenesis is not restricted to the embryonic period. In mammals, milk teeth are replaced by a permanent dentition, and in many vertebrates teeth are replaced continuously throughout life, thus resembling cycling of hair. Although the morphology and molecular regulation of tooth development resembles that of other epithelial appendages, the final outcome of tooth development has very special characteristics. They come in a variety of forms which serve special functions in catching, chopping and chewing food. Because of their masticatory functions, the teeth are necessary for the survival of most animals. The teeth of modern vertebrates are enamel covered structures which also contain two other types of mineralized tissues, dentin and cementum. The enamel matrix is of epithelial origin and is composed of unique enamel proteins. Dentin and cementum are mesenchymal tissues in which the major component is type I collagen. Dentin also contains specific molecular components not found in bone.3 Its structure also differs from bone, as the odontoblasts do not become embedded in the extracellular matrix like the osteoblasts in bone, but remain as a columnar cell layer at the dentin-pulp interface. Each odontoblast leaves one cell process behind it while depositing dentin, and these processes occupy the dentinal tubules characteristic of dentin structure.

Evolution of Teeth Teeth are unique to vertebrates, and during their evolution, a variety of modifications in teeth have been seen.4 The earliest dental-like tissues have been found in more than 500 million year old fossils of the first vertebrates.5,6 These structures contained dentin-like tissue, but they were not restricted to the oral region and formed a dermal skeleton.7 Their evolution is believed to be associated with the appearance of the neural crest.8 The common embryonic primordia for these dermal denticles and the teeth are called odontodes, and, as jaws evolved, the odontodes in the oral cavity were modified to teeth. The dermal skeleton has been mostly lost during evolution, but derivatives of odontodes still persist outside the mouth region in some fish, e.g., in sharks as placoid scales.

The Teeth as Models for Studies on the Development and Evolution of Organs

159

Fig. 9.2. Comparison of the dentition of the common opossum (Didelphys virginiana), a marsupial, and the rat (Mus rattus), a rodent. Variations can be seen in the total number of teeth (opossum 50, rat 16), in the number of teeth in different subgroups (incisors: opossum 9, rat 4; canines: opossum 4, rat 0; premolars: opossum, 12, rat 0; molars: opossum, 16, rat 12) and in the shape of the teeth (e.g., the largest teeth in opossums are the canines, whereas in rats they are the incisors). Reprinted from Osborn HF. Evolution of mammalian molar teeth. London: The Macmillan Company 1907.

Although the basic developmental anatomy of tooth morphogenesis has been conserved to a high degree and the morphological steps of tooth development (Fig. 9.1) are very similar in all living vertebrates, there are a variety of modifications in the ways in which the dentitions are organized and in which they function in different animals. The number of teeth vary, as well as the shape and structure of individual teeth (Fig. 9.2). Birds, for example, have lost all their teeth, and so have turtles. There are also some toothless mammalian species. The mammalian dentition differs in many aspects from those of other vertebrates. In mammals, the teeth form a single row in the mandible and maxilla, whereas in fish there are many rows of teeth and in some, like the zebrafish, they are found in the pharyngeal region. Most fish, reptiles and amphibians have a homodont dentition, meaning that all teeth have the same shape. The teeth have simple conical structures and they are also replaced throughout life, i.e., they are polyphyodont. Mammals have a heterodont dentition, involving four different classes of teeth (Fig. 9.2). Furthermore, the mammalian dentition is replaced only once; they have a primary and secondary dentition. The general dental formula in mammals (number of teeth in different classes in each half of the maxilla and mandible) is three incisors, one canine, four premolars and three molars. The different groups are characterized by their position and shapes. The incisors and canines are located in the anterior of the mouth, and they are morphologically simple, having a cone or blade-like cutting edge. The premolars and molars occupy the more posterior buccal area in the mouth and have a large occlusal surface which is used for grinding food. There is a left-right symmetry in the dentition, but the shapes of individual teeth are asymmetric in different axes (buccal-lingual, anterior-posterior). There are extensive evolutionary modifications in the dental formulae between different mammals. Many mammalian species have lost several teeth. In man, one incisor and one premolar have been lost. The rat, on the other hand, has lost two incisors as well as all canines and premolars, having only one incisor and three molars left in each half of the jaws (Fig. 9.2). In general, the loss of teeth during evolution appears to have happened in the sequence opposite to that in which the teeth develop during ontogenesis. (The teeth in the

160

Molecular Basis of Epithelial Appendage Morphogenesis

different tooth groups develop in an anterior to posterior direction, and the first teeth that have been lost during evolution are the most posterior ones; for example the third incisor and the fourth, and subsequently third, premolars.) In addition to the number of teeth, there are major differences in the shapes of teeth between different mammalian species. The shapes reflect the nature of food the animal eats. The occlusal surfaces of premolars and molars, for example, have varying numbers of cusps which also are of different sizes. In predatory animals the carnassial teeth have a huge cusp which helps in slicing the prey. In many animals the molars have crests connecting the cusps, which increases the efficiency of grinding food. For instance, molars of herbivorous animals are characterized by large surface areas with multiple crests which facilitate the grinding of leaves and hay. In fact, the evolution of mammals has been best documented in their teeth.9,10 In particular, evolutionary changes in the size and location of the cusps and crests in the premolars and molars are correlated with changes in the diet of mammals. For example, the last-forming cusp of the upper molars, called the hypocone, has evolved independently more than 20 times.11 As the hypocone effectively doubles the occlusal area devoted to crushing food, its evolution is related to increase in consumption of plants. Thus the hypocone is observed to have evolved independently at the bases of mammalian radiations leading to specialized herbivory. Moreover, a recent study quantified morphological diversity of fossil ungulate molars, rather than their taxonomic diversity, allowing a new approach for the analysis of adaptive radiations. This study showed that a concomitant increase in the number of crests (shearing blades) on molars with hypocones has happened independently among northern continents.12 A high number of crests is correlated with consumption of fibrous plants, and the acquisition of crests is thus related to the shift towards specialized herbivory, ultimately driven by climatic cooling and drying.

The Neural Crest The mesenchymal component of teeth has its origin in the neural crest, and it is commonly believed that the evolution of the neural crest was critical for the appearance of the teeth.8 That the neural crest cells significantly contribute to tooth development was first demonstrated in amphibians.13 Circumstantial evidence that neural crest cells also contribute to mammalian tooth development was later provided by tissue recombination experiments in which dental epithelium was cultured with mesenchymal cells of different origins. Only neural crest derived cells, including those from the trunk region, took part in tooth development, whereas mesoderm-derived mesenchyme did not.14 Transplantation studies using the quail nuclear marker showed some time ago that the majority of the connective tissue and skeletal elements in the avian jaws derive from the neural crest.15,16 As birds do not have teeth, the neural crest contribution to teeth could unfortunately not be analyzed in these experiments. Recently, labeling of neural crest cells and analysis of their migration has been successful in mouse and rat embryos. Labeling of neural crest cells in three to eight somite mouse and rat embryos, and their follow-up using whole embryo cultures, indicated that specifically the mesenchymal cells beneath the surface ectoderm in the maxillary and mandibular processes are of neural crest origin17-19 (Fig. 9.3A). Direct evidence indicating that dental mesenchyme also derives from the neural crest was recently provided.19 They dissected the mandibular processes of rat embryos after first having labeled the neural crest cells by DiI, and cultured the whole embryos for two days. The jaw explants were allowed to develop for an additional six days in organ culture. Tooth development had reached the bud stage, and labeled cells could be demonstrated in the dental mesenchyme, thus confirming that neural crest cells are actually involved in tooth development (Fig. 9.3B).

The Teeth as Models for Studies on the Development and Evolution of Organs

161

Fig. 9.3. Contribution of neural crest cells to facial processes and dental mesenchyme in rat embryos. (A) Posterior midbrain cells have been labeled at the 3somite stage with DiI. The labeled cells have migrated to the mandibular prominence, the maxillary prominence and the periocular region (lateral view of a wholemount embryo). (B) Posterior midbrain cells have been labeled at the 4-somite stage. In the cultured mandible, labeled cells (arrows) are visible in the dental mesenchyme surrounding the dental epithelium. DE, dental epithelium, DM, dental mesenchyme, E, eye primordium, FNM, frontonasal mass, Md, mandibular prominence, Mx, maxillary prominence, PMB, posterior midbrain, S3, somite stage 3, TG, trigeminal ganglion. Scale bar, 100 mm. Reprinted with permission from Imai H et al, Dev Biol 1996; 176:151-165.

Patterning of Teeth Mammalian teeth are sequentially arranged structures and the different tooth groups, incisors, canines, premolars and molars, show characteristic differences in morphology. Each tooth group forms from one epithelial thickening, the dental lamina, and the development starts with the most anterior tooth and proceeds posteriorly. Two different theories have been put forward to explain the segmental patterning and shape differences amone the teeth. The field theory assumes that concentrations of chemical morphogens regulate different morphogenesis of initially identical primordia.4,20 The field theory has received some support from a study showing differences in concentrations of retinoic acid between the anterior and posterior parts of the jaws.21 According to the other theory, the clone model, the stem cells giving rise to different classes of teeth differ from each other initially.22 Currently, after the discoveries of the Hox-genes, and demonstrations that they code positional identities of cells in the anterio-posterior axis development in Drosophila as well as in vertebrates,23 the clone theory has become more popular. There is circumstantial evidence to support the clone model. The above mentioned studies on neural crest cell migration have shown that the tooth bearing parts of the mandibular and maxillary processes are colonized by neural crest cells from the midbrain region.19,24 Furthermore, the final position of the cells in the maxillary and mandibular processes is determined by the original position of the cells in the neural crest as well as by the time when the cells leave the crest.19 The neural crest cells maintain their patterns of Hox gene expression during migration, and hence it is possible that this array of expressed genes, which has been called the Hox code,25 determines their identity in tooth development. However, it is also possible that the dental identity of the migrating cells is further regulated by

162

Molecular Basis of Epithelial Appendage Morphogenesis

the environment in which they migrate. The cells have been shown to migrate as a sheet along a subectodermal pathway as far as the distal aspect of the branchial arches.18 Hence, it is possible that the neural crest cells have acquired positional identity by the time they reach their final destination in the region of tooth development. Dlx2 expression, for instance, has been correlated with neural crest cell development in the branchial arches.26 It has been suggested that the overlapping expression domains of several homeobox-containing transcription factors, including Dlx and Msx genes in the jaw mesenchyme, would specify the identity of teeth in the incisor, canine, premolar and molar fields.27,28 This theory was recently supported by the interesting finding that double knockout mice deficient for Dlx1 and Dlx2 specifically lack upper molar teeth.29 All incisors, as well as lower molars, developed normally in these mutants. So far one homeobox gene, Barx1, shows tooth typespecific expression (Fig. 9.4A).30 It is present in mouse molar mesenchyme and absent from incisors during morphogenesis. It still remains to be shown whether the segmental identity of teeth is regulated by neural crest derived mesenchymal cells or by the stomodeal ectoderm, and whether this is associated with the initiation of tooth development. In the case of many other ectodermal appendages the mesenchyme determines the position of structures, whereas the epithelium responds according to its inherent capabilities.31 There is also such evidence from interspecies tissue recombination experiments in tooth development. Combination of mouth mesoderm from frog embryos, which normally do not develop teeth, with flank ectoderm from salamanders results in tooth development.32 Heterotopic tissue recombination experiments in mouse embryos have shown that the presumptive dental epithelium instructs tooth morphogenesis prior to the bud stage.14,33 These studies showed that in 9 day old mouse embryos the rostral, but not caudal, epithelium of the mandibular arch is capable of instructing tooth development in trunk neural crest as well as in the mesenchyme of the second branchial arch. However, as the neural crest cells have already reached the branchial arches at the time of the experiments, they may have induced the epithelium to acquire specific tooth forming capacities before the dissection of tissues. Recently, the homeobox-containing transcription factor Otlx2 (RIEG) was shown to be already expressed at embryonic day 8.5, specifically in the rostral stomodeal epithelium, and its expression continued in dental epithelium throughout tooth morphogenesis (Fig. 9.4B).34 This is so far the earliest gene associated with tooth epithelium. However, as it is expressed throughout the dental lamina in the mouse embryo and is not restricted to different dental fields, it appears not to be involved in specification of dental identity. Instead it may be a key regulator of the epithelial response, i.e., tooth formation, to the mesenchymal influence, which perhaps determines the identity of the structures to be formed.

Morphogenesis of Individual Teeth: Regulation by Epithelial-Mesenchymal Interactions We know much more about the molecular mechanisms regulating morphogenesis of single teeth than about those involved in dental patterning and initiation of tooth development. The mechanisms of morphogenetic regulation are largely associated with cellular signaling in the mediation of epithelial-mesenchymal interactions. In fact, the teeth belong to those organs in which the molecular basis of epithelial-mesenchymal signaling has been elucidated to a significant extent during recent years.35 It has become clear that these molecular mechanisms have been conserved to an astonishing extent during evolution, and that the same signals apparently regulate morphogenesis in all vertebrate organs, including epithelial appendages.

The Teeth as Models for Studies on the Development and Evolution of Organs

163

Fig. 9.4. Expression of Barx1 and Otlx2 in newborn mouse molars and incisors. (A) Barx1 is expressed in the mesenchyme of the first, second and third molar, but not in the incisor. (B) Otlx2 is expressed in the epithelium of the first, second and third molar and also in the incisor epithelium. 1m, first molar; 2m, second molar; 3m, third molar; i, incisor. Reprinted with permission from Mucchielli M et al, Dev Biol 1997; 189: 275-284.34

The morphogenesis of all epithelial appendages is regulated by a sequence of reciprocal interactions between the epithelial and mesenchymal tissue components. This has been demonstrated in classic tissue recombination experiments. The significance of epithelialmesenchymal interactions for tooth development was first demonstrated using amphibian tissues,13,32 but subsequently mostly rodent tissues have been analyzed. As mentioned above, the presumptive dental epithelium governs tooth morphogenesis prior to the bud stage.14,33 The potential to instruct tooth morphogenesis, however, shifts to the dental mesenchyme during budding and, as demonstrated by Kollar and his associates,36,37 the dental papilla of cap and bell stage teeth regulates tooth shape development, i.e., a molar tooth develops when molar mesenchyme is cultured with incisor epithelium and vice versa. Furthermore, the dental mesenchyme is able to instruct the differentiation of ameloblasts and enamel secretion in nondental epithelium.37 Hence, in addition to morphogenesis, cell differentiation is also regulated by epithelial-mesenchymal interactions. During the last 20 years numerous molecules have been associated with tissue interactions during tooth morphogenesis. (A database describing some expression changes during tooth development is available at the Internet location http:/honeybee.helsinki.fi/toothexp.) Components of the basement membrane separating the epithelium and mesenchyme have been shown to be important for the terminal cell differentiation of odontoblasts.38,39 It was proposed that the extracellular matrix molecule fibronectin functions in interactions between the basement membrane and the cell surface of the differentiating odontoblast.40 More recently it has been shown that signal molecules in the BMP family (see below) have the capacity to trigger odontoblast differentiation, but this requires matrix molecules in

164

Molecular Basis of Epithelial Appendage Morphogenesis

addition.41 It is possible that the differentiation of odontoblasts requires a close association and molecular interactions with the basement membrane and, in addition, soluble signals which may be bound to the basement membrane. The extracellular matrix glycoprotein tenascin-C and the cell surface proteoglycan syndecan-1 were among the first molecules that were shown to be associated with and regulated by epithelial-mesenchymal signaling during early tooth morphogenesis.42-44 TenascinC and syndecan-1 were shown to interact with each other and, as they are expressed by organ-specific mesenchyme in several other organs as well, it was suggested that they may regulate the condensation of mesenchymal cells.45 Tenascin-C and syndecan-1 are both members of gene families, but the exact roles of these molecules are still unknown. Tooth development occurs normally in tenascin deficient mice.46 On the other hand, as syndecan binds heparin-binding growth factors such as FGFs, it may be involved in cell-growth factor interactions.47 Numerous signal molecules and growth factors in several different families, as well as their specific receptors, have been associated with epithelial-mesenchymal signaling during tooth morphogenesis (Figs. 9.5 and 9.6).35,48 In situ hybridization analysis of the patterns of expression of individual signals and their receptors indicate that in some cases they are restricted to either epithelial or mesenchymal tissues and in some cases to a certain developmental stage. However, in many cases the signals seem to be used several times during morphogenesis and/or they appear to transfer messages in both directions between the interacting tissues. The effects of different signals have been experimentally analyzed in organ culture studies.49 Recombinant signal proteins have been applied to isolated dental tissues with agarose or heparin acrylic beads to mimic the effects of epithelium on mesenchyme or vice versa. The cellular responses have been monitored by different methods, e.g., by analyzing cell proliferation with BrdU incorporation or gene expression by in situ hybridization. In such studies it has been possible to elucidate the signaling networks involved in epithelial-mesenchymal interactions.50-52 Bone morphogenetic proteins (BMPs) were the first signals that were identified in the transmission of inductive interactions between the dental epithelium and mesenchyme.51 BMPs constitute one of the signal families which has widespread signaling functions throughout the animal kingdom.53 The best characterized member is the Drosophila morphogen Dpp, which is homologous with the vertebrate BMP-2 and BMP-4. We showed that BMP-2 and BMP-4 recombinant proteins mimicked the effect of presumptive dental epithelium on mesenchyme at the time when the instructive capacity is known to reside in the epithelium (see below). Analysis of the expression patterns of different Bmps indicates that, in addition to Bmp-2 and -4, Bmp-7 is also expressed early in dental epithelium (Fig. 9.5). These three Bmps shift between epithelium and mesenchyme during advancing morphogenesis and they are also expressed in the enamel knot (see below). In addition they are associated with the differentiation of odontoblasts and ameloblasts.41,54 Bmp-5 expression, on the other hand, is confined to the ameloblasts.54 Several of the serine-threonine kinase receptors for BMPs are expressed during morphogenesis but the patterns do not appear to be as clearly restricted spatiotemporally as those of the ligands (ref. 55 and Åberg et al, unpublished observations). In addition to Bmps, other members of the TGF-β superfamily, notably TGF-β-1, -2 and -3 are also present in teeth and may be associated with morphogenesis.56,57 Several of the fibroblast growth factor (FGF) family members have been associated with tooth morphogenesis. Fgf-3 (int-2) was the first to be localized.58 Its expression is confined to dental papilla mesenchyme and it is downregulated with advancing morphogenesis (our unpublished observations). Fgf-4, -8 and -9, on the other hand, are expressed exclusively

The Teeth as Models for Studies on the Development and Evolution of Organs

Fig. 9.5. Bone morphogenetic proteins (Bmp) are expressed in the epithelium during early tooth development. Mouse mandibular molar tooth germs. (A) Bmp-2, early E12. (B) Bmp-4, E11. (C) Bmp-7, late E12. (Courtesy Thomas Åberg).

Fig. 9.6. Fibroblast growth factor-8 (Fgf-8) is expressed in the epithelium during initiation of tooth development. Frontal section through the head of an E11 mouse embryo. In situ hybridization analysis shows intense Fgf-8 expression in the thickened oral epithelium at the sites of developing maxillary and mandibular molar teeth. (Courtesy Päivi Kettunen).

165

166

Molecular Basis of Epithelial Appendage Morphogenesis

in dental epithelial cells.50,59,60 Fgf-8 is only present in the presumptive dental epithelium prior to budding (Fig. 9.6), and Fgf-4 is restricted to the enamel knots (Figs. 9.7-9.9, see below). Fgf-9 expression appears to cover the domains of both Fgf-4 and -8 and in addition it is widely expressed in the dental epithelium during the bell stage, in association with the terminal differentiation of the odontoblasts and ameloblasts. Their respective tyrosine kinase receptors are present in both epithelial and mesenchymal tissues in the tooth.61,62 Of the hedgehog family, Sonic hedgehog (Shh) is expressed in dental epithelium at several stages, starting in the early epithelial thickenings and then reappearing in the enamel knot (Fig. 9.8, see below) and subsequently it is expressed in the ameloblast cell lineage.63-65 Its receptor Patched, which is a multipass membrane protein,66 is expressed widely in the dental mesenchyme (our unpublished observations). Other conserved signals that have been analyzed during tooth morphogenesis include epidermal growth factor (EGF). EGF receptor is expressed both in dental epithelium and mesenchyme and is associated and regulated by tissue interactions.67 However, we have been unable to detect expression of either Egf or Tgf-α, another EGF receptor ligand in teeth, by in situ hybridization (Partanen et al, unpublished observations). Egf transcripts have been detected by PCR methods.68 Surprisingly, no dental anomalies were reported in EGF receptor deficient mice although they had major defects in development of several other epithelial appendages.69,70 The Notch receptor as well as its ligand Serrate (Jagged) are also expressed during several stages of tooth morphogenesis, and both in epithelial and mesenchymal tissues.52,71 Interestingly, Notch-1,-2 and -3 genes are already downregulated in the dental epithelium at the time of dental lamina formation. Throughout subsequent tooth morphogenesis the cells of the ameloblast cell lineage, i.e., inner enamel epithelial cells, do not express the Notch genes, whereas all other dental epithelial cells express them intensely. In Drosophila the expression of the Notch gene is associated with cell fate specification72 and hence it is possible that Notch genes function in the determination of the dental epithelial cells. Furthermore, Notch and Serrate genes are regulated by tissue interactions as well as by BMPs and FGFs in dental tissues, thus suggesting that signaling through this pathway is linked with other signaling networks.52,71 In vitro experiments, in which the expression of hepatocyte growth factor (HGF, scatter factor) was inhibited by antisense oligonucleotides, suggest that this signal is needed for tooth morphogenesis. HGF is expressed in dental mesenchyme and its receptor c-Met in epithelium,73 and the prevention of HGF expression inhibited the progression of morphogenesis from cap to bell stage.74 In similar antisense experiments the prevention of expression of the precursor of substance P, a tachykinin neurotransmitter, resulted in inhibition of tooth as well as mammary gland morphogenesis.75 Interestingly, neurotrophins and neurotrophin receptors (both the low affinity receptor, LANR, and tyrosine kinase trk-receptors) are also expressed during tooth morphogenesis. Although their expression is largely linked with the development of innervation, some expression domains show no relation to nerve growth but are rather associated with epithelial-mesenchymal interactions.76,77

The Enamel Knot as a Signaling Center The enamel knots are transient clusters of dental epithelial cells which were observed many decades ago in cap stage teeth.78,79 They were subsequently largely neglected and sometimes even viewed as histological artefacts. Recent work from our laboratory has led to the acceptance of the enamel knots as real structures and to suggestions that they may represent signaling or organizing centers, perhaps regulating tooth shape.59,64 The enamel knots are first seen at the tips of the tooth buds just prior to their development into cap stage. A few cells of the epithelium stop proliferating, as seen by the lack of

The Teeth as Models for Studies on the Development and Evolution of Organs

Fig. 9.7. Fgf-4 expression is restricted to the enamel knot. (A) The enamel knot can be seen as a cluster of cells in the middle of the epithelium in a cap-stage E14 molar. (B) Fgf-4 expression in the enamel knot.64

BrdU

Shh

Bmp-4

Fgf-4

13 d

distal lingual

14 d

buccal

anterior

200 µg

Fig. 9.8. The enamel knot is a signaling center. Three-dimensional reconstructions of E13 (upper row) bud stage and E14 (lower row) cap-stage teeth from BrdU labeled or in situ hybridized serial sections (as in Figs 5C and 6). The light lines represent the epithelial-mesenchymal interphase. BrdU labeling shows a smaller (E13) or a larger (E14) area of nonproliferating cells, which specifies the extent of the enamel knot. Shh, Bmp-4 and Fgf-4 expression overlaps with the enamel knot.64

167

Molecular Basis of Epithelial Appendage Morphogenesis

168

BrdU

Fgf-4

Fig. 9.9. Localization of secondary enamel knots. Three-dimensional reconstruction of E16 bell-stage tooth from BrdU labeled or Fgf-4 hybridized serial sections. The lines represent the epithelial-mesenchymal interphase. BrdU labeling shows cell proliferation in all areas except at the sites of the secondary enamel knots (marked with arrows). The nonproliferating sites also express Fgf-4.59

BrdU incorporation. The enamel knot grows as the bud transforms to a cap, at which stage it is readily visible in histological sections as a tightly packed epithelial cell aggregate. The appearance and shape of the enamel knot has been analyzed in detail in the first mandibular molar tooth in mouse embryos by three-dimensional computer based reconstruction of serial sections (Fig. 9.8). The analysis was based on BrdU labeling, and the extent of the enamel knot was determined as the region of nonproliferating cells. The enamel knot was shown to be a torpedo-shaped structure, the development of which started in the mesial part of the tooth germ at the late bud stage, and progressed distally.59 The assumption that the enamel knot represents a specific cell lineage with a unique fate is supported by expression of the cyclin dependent kinase inhibitor p21, which is associated with terminal differentiation of many cell types.80 We have shown that p21 expression is stimulated by BMP-4 protein and have suggested that BMP-4, which is intensely expressed in the dental mesenchyme during the bud stage, might be involved in the induction of enamel knot formation.81 The first indication that the enamel knot may have a signaling function came from in situ hybridization studies in which Fgf-4 gene expression was localized exclusively in the enamel knot (Fig. 9.7).59,82 Subsequently several other signals have been shown to be expressed in the enamel knot. These include Bmp-2, -4 and -7, and Shh64 as well as Fgf-9 (Fig. 9.8).50 These signals show nested expression patterns and differences in the onset and termination of expression. Fgf-4 expression correlates rather well with the nonmitotic cells, although the expression domain is a few cell layers more restricted and starts slightly later.59 Shh, Bmp-2 and Bmp-7 are already present in budding epithelium prior to the exit of the distal tip cells from the cell cycle. Bmp-4, on the other hand, starts to be expressed during advanced cap stage starting from the distal region of the knot (Fig. 9.8).81 Although there is no direct evidence for an organizing capacity of the enamel knot, this theory is supported by the fact that the same signals are expressed in well known organizing centers in the embryo. In the developing limb the zone of polarizing activity (ZPA), which regulates anteroposterior patterning, expresses Shh83 and the apical ectodermal ridge (AER), which controls proximodistal growth in the limb and interacts with the ZPA, expresses Fgf-4.82 In addition, both ZPA and AER express the same Bmps as the enamel knot.84 The notochord, which regulates patterning of the neural tube and somites, also expresses Shh, and

The Teeth as Models for Studies on the Development and Evolution of Organs

169

the ectoderm overlying the neural tube, which participates in this patterning, expresses Bmp-4 and Bmp-7.85-87 Shh expression, as well as Bmps and FGFs, have also been localized to budding epithelium during hair and feather development and in the lungs,88-91 suggesting conserved actions during epithelial appendage development. However, no morphologically distinct epithelial structures resembling the enamel knot have been described in the other epithelial appendages. An interesting feature of the enamel knot is that its cells undergo apoptosis.92 It starts to disappear during cap stage, starting from its distal end.81 The removal therefore occurs in the opposite direction than that in which it is formed. Consequently, no enamel knot is seen at late cap stage. We have proposed that apoptosis may be a general mechanism whereby the function of signaling centers may be extinguished.92 Cell death is apparent also in the AER92,93 and ZPA, as well as notochord, as they are removed with advancing development. The expression pattern of Bmp-4 in the enamel knot shows striking colocalization with apoptosis. As BMP-4 regulates programmed cell death during embryonic development in the rhombomeres and in the interdigital mesenchyme,94,95 it is possible that this signal may have a similar function in the enamel knot. We have proposed that the enamel knot determines the site of the first cusp of teeth and regulates the formation of other cusps in molar teeth. There is an enamel knot also in the incisors during cap stage, and in molars new enamel knots appear at the sites of new cusps.59 These secondary enamel knots also express Fgf-4 and their cells do not divide (Fig. 9.9). We have proposed that FGF-4 may regulate the growth of the cusps by stimulation of cell proliferation. Receptors for FGFs are present both in the mesenchyme and epithelium around the enamel knot,62 and FGF-4 stimulates the proliferation of both cell types.59 Hence, the spatial pattern of the enamel knots would determine the sites of cusps and their timing would regulate the onset of cusp formation. Because of the way that tooth develops, the cusp that starts to form first will become the highest, and the subsequently forming cusps will end up being progressively shallower. Hence the timing of initiation of cusp formation determines its relative height as compared to other cusps.10,59 The function of FGF-4 and FGF-9 may be to promote cuspal growth. The roles of the different signals in the enamel knot are not understood at present. It is possible that some molecules act as autocrine signals within the enamel knot cells. Some may act as planar signals within the dental epithelium, whereas the underlying dental papilla mesenchyme presumably is the target of some signals. Patched, the Shh receptor, is expressed in the dental papilla mesenchyme, suggesting that the mesenchyme may be the target for the action of Shh (our unpublished observations). Expression of Shh, as well as of Bmp-2 and Bmp-7, appears to spread from the enamel knot along the enamel epithelium during late cap stage, and they cover the coronal part of the dental epithelium during subsequent stages. Whether this spreading is associated with differentiation of the odontoblasts and/or ameloblasts or, perhaps, with development of cusp morphology remains to be shown.

Signaling Networks and Transcription Factors The signaling cascades regulating tooth morphogenesis, as well as the development of other organs, should be seen as a continuous discussion between cells and tissues that directs the progression of development. It has become evident that the signaling networks are complex and also include a multitude of feed-back regulatory loops in which different signaling pathways are connected to each other. In general, when a signal acts on the responding cell it binds to a specific cell surface receptor and this leads to a multistep intracellular process that results in regulation of transcription in the nucleus. This leads to changes in various cellular functions such as the cell cycle or adhesiveness of the cell, and importantly the cellular responses also include the continuation of the signaling cascade, i.e., the expression

170

Molecular Basis of Epithelial Appendage Morphogenesis

of the next reciprocal signals as well as the next receptors that the cell needs to respond to subsequent signals.96 Some transcription factors have been identified which are involved in the epithelialmesenchymal signaling during tooth morphogenesis, and their roles in the signaling cascades are starting to be elucidated. In the dental mesenchyme Msx-1 appears to act both downstream and upstream of BMP signaling. Msx-1 is regulated by epithelial signals during budding of the tooth germ, and BMP-2 and -4 mimic this (Fig. 9.10).51 In addition, several FGFs, including FGF-2, -4, -8, and -9, upregulate Msx-1 expression in the dental mesenchyme.50,97 Interestingly, Msx-1 deficient mice have no teeth; more precisely, development is halted at bud stage.98 Analysis of these mutant teeth has shown that Bmp-4 expression in mesenchyme is not upregulated as in wild type animals, indicating that Msx-1 also acts upstream of the BMP signal.97 This was further supported by the rescue experiment in which the bud stage tooth germs dissected from the Msx-1 deficient embryos were shown to reach cap stage when cultured in the presence of BMP-4 protein (for more details, see chapter 14). Another transcription factor which is associated with the transmission of epithelialmesenchymal signaling is the HMG domain factor Lef1. Like the Msx-1 knockouts, Lef1 knockouts also lack teeth, and the development is arrested at bud stage.99 Also, hair and mammary gland morphogenesis is inhibited in these mice, indicating once again the conservation of molecular mechanisms of organogenesis. Overexpression of Lef1 in epithelial cells in transgenic mice resulted in increased invagination of epithelium and in formation of extra hair follicles and a tooth-like structure.100 Elegant tissue recombination experiments using tissues from Lef1 deficient and wild type mouse embryos showed that, although Lef1 is expressed throughout tooth morphogenesis and its expression is not restricted to either epithelial or mesenchymal tissues, it is needed only in epithelium and only during early development.101 Normal teeth developed from explants which consisted of exclusively mutant tissues—both epithelium and mesenchyme, if the mesenchyme had first been exposed to wild type epithelium before bud stage. Hence, Lef1 appears to be involved in the regulation of an epithelial signal acting on dental mesenchyme during the bud stage of tooth morphogenesis. It is not clear which signaling pathway Lef1 is associated with, but it does not appear to be involved in the same pathway as BMP and Msx-1 in tooth morphogenesis.97,101 Lef1, as well as its Drosophila homologue pangolin, have been recently associated with the Wnt/wingless pathway.102,103 A third transcription factor that was recently identified as part of epithelial-mesenchymal signaling in tooth development is Pax9. It is a member of the paired box-containing transcription factor genes. The tooth phenotype of Pax9 knockouts is similar to those of Msx-1 and Lef1 knockouts, i.e., development is arrested at the bud stage.104 Like Msx-1, Pax9 is also expressed intensely in the dental mesenchyme, and tissue recombination studies showed that it is regulated by the dental epithelium.104a Interestingly, in the early mouse mandibles Pax9 is stimulated by the rostral epithelium but not by the caudal epithelium, which is in line with the earlier observations that only rostral epithelium has the capacity to govern tooth morphogenesis.14 FGF-8 which is expressed in this epithelium at the early stages of tooth morphogenesis, was shown to stimulate Pax9 expression. It is noteworthy that tooth development is arrested in all three transcription factor knockouts at the bud stage. The transition from the bud to cap stage involves the invagination of the undersurface of the epithelial bud and the formation of the dental papilla in the adjacent mesenchyme. We have speculated that the arrest in the development of the knockout teeth may be associated with the lack of the formation of the enamel knot.81 The induction of the enamel knot may be a key event in tooth morphogenesis, and it is possible that proper enamel knot formation and maintenance requires the coordination of several signaling networks.

The Teeth as Models for Studies on the Development and Evolution of Organs

171

Fig. 9.10. BMP-2 induces Msx-1 expression in the dental mesenchyme. (A) A BMP-2 coated agarose bead was placed on an isolated E12 mouse dental mesenchyme and cultured for 24 h. After culture in situ hybridization was performed with a Msx-1 antisense probe. (B) No expression is seen using a Msx-1 sense probe. (Courtesy Carin Sahlberg).

Disrupted Tooth Development Tooth development is obviously under strict genetic control and hence it is apparent that disruptions of tooth development resulting in missing teeth are caused by genetic rather than environmental influences. Loss of teeth is a frequent phenomenon during evolution of animal species, and one or more missing teeth is a rather common, mostly inherited condition in human beings. There are also some natural mutations in animals in which some teeth are missing and, as described above, defective tooth development has been caused by targeted mutations of several genes in transgenic mice. Knowing the sequential nature of regulative events during tooth development and the complexity of the signaling networks mediating epithelial-mesenchymal interactions, it is conceivable that loss of teeth can result from defects in many different genes and that such genes may be associated with inductive signaling. Furthermore, it is apparent that development may be interrupted at different stages of tooth morphogenesis. These assumptions are supported by the phenotypes of the transgenic mice that have been so far produced and show abnormal tooth morphogenesis. First, all such targeted genes are associated with epithelial-mesenchymal signaling. Targeted disruption of activin bA or follistatin results in failure of incisor development, whereas molars reach abnormal looking bell stages.105 The three lines of knockout mice in which the Msx-1, Lef1, and Pax9 transcription factors were targeted and tooth development was arrested at bud stage in all cases,98,99,104 show that tooth development can be blocked at the same stage by disruption of different genes. In Patch mice, which are spontaneous platelet-derived growth factor receptor alpha subunit (PDGFRα) mutants, the invagination of the dental epithelium occurs, but there is incomplete condensation of the dental mesenchyme.106 The molecular mechanisms of the loss of teeth during evolution has recently been approached by analysis of the vestigial tooth buds that are present in many animals at sites where teeth are missing. In mice, which have lost two incisors as well as all canines and premolars, rudimentary tooth buds are present in the maxilla in the diastema region (the toothless region between the incisor and the first molar).107 When the expression of several genes associated with early tooth development was analyzed, it was observed that Msx-2

172

Molecular Basis of Epithelial Appendage Morphogenesis

was not upregulated in the dental epithelium.108 More recent work from our laboratory indicates that in vestigial tooth buds in voles, development is arrested prior to the formation of the enamel knot, and that the expression of Shh, Msx-2, p21 and Fgf-4 fail to appear in the dental epithelium.109 The actual genetic causes for the arrested tooth development in vestigial tooth anlagen remain to be unraveled. In cases of hypodontia, i.e., conditions where one or more teeth are missing, the remaining teeth commonly show reduction in size and conical shapes. This is seen in the most common condition of hypodontia in man, the so-called simple hypodontia, where one or more lateral incisors or second premolars are missing. Peg shaped lateral incisors or small premolars are frequent in these cases. It is noteworthy that the teeth that are missing are the last that develop in their respective tooth groups. Hence this is analogous to the evolutionary reduction in tooth number, which has also happened in the sequence opposite to that of tooth formation. Simple hypodontia is autosomally dominantly inherited, but the causative gene(s) have not been identified. Several candidate genes, all involved in signaling pathways, have been excluded, including the Msx-1 and Msx-2 genes, as well as Egf, its receptor and Fgf-3.110,111 Interestingly, a mutation in the homeodomain of the Msx-1 gene was recently shown to cause oligodontia, a rare condition involving lack of eight or more teeth, in one family.112 The gene causing anhydrotic ectodermal dysplasia (EDA), the most common form of ectodermal dysplasias, was cloned recently113 (see also chapter 21). This X-chromosomal gene encodes a novel cell surface molecule, but its function is so far unknown. EDA is characterized by defective development of epithelial appendages, notably teeth, hair and sweat glands. The tooth phenotype varies, extending from complete lack of teeth, or anodontia, to a few missing teeth, and the remaining teeth show aberrant development such as conical shapes. Tabby is a natural mouse mutation that has long been believed to be the mouse homologue of human EDA. Teeth, hair and a multitude of exocrine glands are affected in the Tabby mouse and the phenotype varies according to the mouse strain.114-116 The recent cloning of the Tabby gene has verified that it is the EDA homologue and shown that there are alternatively spliced forms (Srivastava et al116a). In situ hybridization analysis of Tabby expression indicates that it is expressed by epithelial cells in the developing skin and teeth. Interestingly, the expression is intense in the dental epithelium at cap and bell stages in the outer enamel epithelium (Fig. 9.11). Although the function of the EDA/Tabby gene is not understood at present, it can be speculated that it too is associated with epithelial mesenchymal signaling. It is possible that it is associated with the EGF signaling pathway, because EGF injections to newborn mice partially rescue the sweat gland phenotype in Tabby mice.117 Three other mouse mutants are known which have a similar phenotype to Tabby: the autosomal recessive mutants crinkled and downless, and the autosomal dominant Sleek.118-120 It is conceivable that their respective genes function in the same molecular pathway as Tabby. Another gene that is presently known to be necessary for normal tooth development in man is RIEG, the gene involved in Rieger syndrome. This is an autosomal dominant disorder that in addition to hypodontia includes anomalies in the eye and umbilicus. The cloning of this gene showed that it is a new homeobox gene, and the analysis of expression of the mouse homologue indicated that it is expressed in the mandibular and maxillary epithelium during early tooth morphogenesis.121 The mouse gene was also cloned independently by another group and called Otlx2 (Fig. 9.4B). It was shown that the expression in the presumptive tooth epithelium starts very early, at E8.5, and that the expression persists in the dental epithelium throughout tooth morphogenesis.34 It is possible that Otlx2 is involved in the initiation of tooth formation. It appears to be associated with epithelial-mesenchymal interactions, and it has been shown that its expression can be induced in nondental, Otlx2 negative epithelium by dental mesenchyme.

The Teeth as Models for Studies on the Development and Evolution of Organs

173

Fig. 9.11. Expression of Tabby in an E15 cap-stage mouse molar is localized to the outer enamel epithelium. (A) Bright-field image with overlay of expression. (B) Dark-field image.

Concluding Remarks Teeth belong to those epithelial appendages which have been actively studied for centuries. The morphological features of their development at the macroscopical and microscopical levels have been described in detail, and they were among the organs that were analyzed by the early experimental embryologists during the first part of this century. The regulatory roles of epithelial-mesenchymal interactions, as well as the neural crest origin of the dental mesenchyme, were first demonstrated in these early experiments using amphibian tissues. Teeth were subsequently among the mammalian organs in which the sequential and reciprocal characteristics of tissue interactions were analyzed during the second half of this century. This knowledge has formed the basis for the more recent studies on developmental regulation at the molecular level. As a result, the tooth is at present one of the vertebrate organs in which the signaling networks have been elucidated to a reasonable extent. Hence, teeth are at present popular subjects in studies on genetic regulation of development, and it can be expected that this information will accumulate with increasing speed. Although the early developmental anatomy of teeth resembles that of other epithelialmesenchymal organs, and the genetic networks regulating morphogenesis are strikingly similar, the teeth are in many ways very special epithelial appendages. Certain unique features in teeth make them exceptionally good objects for studies on developmental as well as evolutionary mechanisms. Their characteristic sequential arrangement in the mouth, the patterning of the various tooth groups, and the origin of the mesenchymal component of teeth in the midbrain neural crest provide excellent models for testing hypotheses on the development of positional identities of cells and cell groups. The different tooth groups with differing forms, such as incisors and molars, provide unique models for the analysis of the process of epithelial morphogenesis and the development of shape. A particularly advantageous feature of teeth is that they are the only epithelial appendages that have been preserved in fossil records. This has allowed detailed descriptions of dental evolution. Morphological evolution obviously is a result of alterations in developmental programs. Hence, teeth should provide a unique tool for the successful combination of developmental and evolutionary information, which could lead to identification of genes and morphogenetic programs responsible for evolution.

174

Molecular Basis of Epithelial Appendage Morphogenesis

References 1. Ten Cate AR. Oral Histology. St Louis: Mosby-Year Book, 1994. 2. Thesleff I, Vaahtokari A, Partanen AM. Regulation of organogenesis. Common molecular mechanisms regulating the development of teeth and other organs. Int J Dev Biol 1995; 39:35-50. 3. Butler. Dentin matrix proteins and dentinogenesis. Connect Tissue Res 1995; 33:59-65. 4. Butler PM. Ontogenetic aspects of dental evolution. Int J Dev Biol 1995; 39:25-34. 5. Smith MM, Hall BK. Development and evolutionary origins of vertebrate skeletogenic and odontogenic tissues. Bio Rev Camb Philos Soc 1990; 65:277-373. 6. Sansom IJ, Smith MP, Smith MM. Dentine in conodonts. Nature 1994; 368:591. 7. Smith MP, Sansom IJ, Repetski JE. Histology of the first fish. Nature 1996; 380:702-704. 8. Smith MM, Hall BK. A developmental model for evolution of the vertebrate exoskeleton and teeth: The role of cranial and trunk neural crest. Evol Biol 1993; 27:387-448. 8a. Osborn HF. Evolution of mammalian molar teeth. London: The Macmillan Company, 1907. 9. Fortelius M. Ungulate cheek teeth: Developmental, functional, and evolutionary interrelations. Acta Zool Fennica 1985; 180:1-76. 10. Jernvall J. Mammalian molar cusp patterns: Developmental mechanisms of diversity. Acta Zool Fennica 1995; 198:1-61. 11. Hunter JP, Jernvall J. The hypocone as a key innovation in mammalian evolution. Proc Natl Acad Sci USA 1995; 92:10718-10722. 12. Jernvall J, Hunter JP, Fortelius M. Molar tooth diversity, disparity, and ecology in Cenozoic ungulate radiations. Science 1996; 274:1489-1492. 13. Sellman S. Some experiments on the determination of the larval teeth in Ambystoma mexicanum. Odontol Tidsk 1946; 54:1-128. 14. Lumsden AG. Spatial organization of the epithelium and the role of neural crest cells in the initiation of the mammalian tooth germ. Development 1988; 103 Suppl:155-169. 15. Noden DM. Craniofacial development: New views on old problems. Anat Rec 1984; 208:1-13. 16. Le Douarin NM, Ziller C, Couly GF. Patterning of neural crest derivatives in the avian embryo: in vivo and in vitro studies. Dev Biol 1993; 159:24-49. 17. Osumi-Yamashita N, Ninomiya Y, Doi H et al. The contribution of both forebrain and midbrain crest cells to the mesenchyme in the frontonasal mass of mouse embryos. Dev Biol 1994; 164:409-419. 18. Trainor PA, Tam PP. Cranial paraxial mesoderm and neural crest cells of the mouse embryo: Co-distribution in the craniofacial mesenchyme but distinct segregation in branchial arches. Development 1995; 121:2569-2582. 19. Imai H, Osumi-Yamashita N, Ninomiya Y et al. Contribution of early-emigrating midbrain crest cells to the dental mesenchyme of mandibular molar teeth in rat embryos. Dev Biol 1996; 176:151-165. 20. Butler PM. Studies of the mammalian dentition. Differentiation of the postcanine dentition. Proc Zool Soc Lond 1939; 109B:329-356. 21. Kronmiller JE. Beeman CS. Spatial distribution of endogenous retinoids in the murine embryonic mandible. Arch Oral Biol 1994; 39:1071-1078. 22. Osborn, JW. Morphogenetic gradients: fields versus clones. In Development, Function and Evolution of Teeth. Butler PM, Joysey KA, eds. New York: Academic Press, 1978:171-201. 23. Hunt P, Krumlauf R. Hox codes and positional specification in vertebrate embryonic axes. Ann Rev Cell Biol 1992; 8:227-256. 24. Köntges, G, Lumsden A. Rhombencephalic neural crest segmentation is preserved throughout craniofacial ontogeny. Development 1996; 122:3229-3242. 25. Hunt P, Gulisano M, Cook M et al. A distinct Hox code for the branchial region of the vertebrate head. Nature 1991; 353:861-864. 26. Robinson GW, Mahon KA. Differential and overlapping expression domains of Dlx2 and Dlx3 suggest distinct roles for Distal-less homeobox genes in craniofacial development. Mech Dev 1994; 48:199-215. 27. Sharpe PT. Homeobox genes and orofacial development. Connect Tissue Res 1995; 32:17-25.

The Teeth as Models for Studies on the Development and Evolution of Organs

175

28. Weiss KM, Ruddle FH, Bollekens J. Dlx and other homeobox genes in the morphological development of the dentition. Connect Tissue Res 1995; 32:35-40. 29. Thomas BL, Tucker AS, Qiu M, Ferguson C, Hardcastle Z, Rubenstein JL, Sharpe PT. Role of DIX-1 and DIX-2 genes in patterning of the murine dentition. Development 1997; 124:4811-4818. 30. Tissier-Seta JP, Mucchielli Ml, Mark M et al. Barx1, a new mouse homeodomain transcription factor expressed in cranio-facial ectomesenchyme and the stomach. Mech Dev 1995; 51:3-15. 31. Sengel P. Morphogenesis of Skin. Cambridge: Cambridge University Press, 1997. 32. Wagner G. Untersuchungen an Bombinator-Triton-Chimaeren. Das Skelett Larvaler Triton-Köphe mit Bombinator-Mesektoderm. Wilhelm Roux Arch. Entw Mech Org 1959; 151:136-158. 33. Mina M, Kollar EJ. The induction of odontogenesis in non-dental mesenchyme combined with early murine mandibular arch epithelium. Arch Oral Biol 1987; 32:123-127. 34. Mucchielli M, Mitsiadis T, Raffo S et al. Mouse Otlx2/RIEG expression in the odontogenic epithelium precedes tooth initiation and requires mesenchyme-derived signals for its maintenance. Dev Biol 1997; 189:275-284. 35. Thesleff I, Nieminen P. Tooth morphogenesis and cell differentiation. Curr Opin Cell Biol 1996; 8:844-850. 36. Kollar EJ, Baird GR. The influence of the dental papilla on the development of tooth shape in embryonic mouse tooth germs. J Embryol exp Morph 1969;131-148. 37. Kollar EJ, Baird GR. Tissue interactions in embryonic mouse tooth germs. II. The inductive role of the dental papilla. J Embryol exp Morph 1970; 24:173-186. 38. Thesleff I, Barrach HJ, Foidart JM et al. Changes in the distribution of type IV collagen, laminin, proteoglycan, and fibronectin during mouse tooth development. Dev Biol 1981; 81:182-192. 39. Ruch JV, Lesot H, Begue-Kirn C. Odontoblast differentiation. Int J Dev Biol 1995; 39:51-68. 40. Lesot H, Kubler MD, Fausser JL et al. A 165 kDa membrane antigen mediating fibronectinvinculin interaction is involved in murine odontoblast differentiation. Differentiation 1990; 44:25-35. 41. Begue-Kirn C, Smith AJ, Ruch JV. Effects of dentin proteins, transforming growth factor beta 1 (TGF beta 1) and bone morphogenetic protein 2 (BMP2) on the differentiation of odontoblast in vitro. Int J Dev Biol 1992; 36:491-503. 42. Thesleff I, Mackie E, Vainio S et al. Changes in the distribution of tenascin during tooth development. Development 1987; 101:289-296. 43. Thesleff I, Jalkanen M, Vainio S et al. Cell surface proteoglycan expression correlates with epithelial-mesenchymal interaction during tooth morphogenesis. Dev Biol 1988; 129:565-572. 44. Vainio S, Jalkanen M, Thesleff I. Syndecan and tenascin expression is induced by epithelial- mesenchymal interactions in embryonic tooth mesenchyme. J Cell Biol 1989; 108:1945-1953. 45. Salmivirta M, Elenius K, Vainio S et al. Syndecan from embryonic tooth mesenchyme binds tenascin. J Biol Chem 1991; 266:7733-7739. 46. Saga Y, Yagi T, Ikawa Y et al. Mice develop normally without tenascin. Genes Dev 1992; 6:1821-1831. 47. Salmivirta M, Heino J, Jalkanen M. Basic fibroblast growth factor-syndecan complex at cell surface or immobilized to matrix promotes cell growth. J Biol Chem 1992; 267:17606-17610. 48. Thesleff I, Sahlberg C. Growth factors as inductive signals regulating tooth morphogenesis. Semin Cell Dev Biol 1996; 7:185-193. 49. Thesleff I, Sahlberg C. Organ culture in the analysis of tissue interactions. In: Mason I, Sharpe P, eds. Methods in Molecular Biology. In Press. 1997. 50. Kettunen P, Thesleff I. Expression and function of FGFs-4, -8 and -9 suggest functional redundancy and repetitive use as epithelial signals during tooth morphogenesis. Dev Dyn 1998; 211:256-268.

176

Molecular Basis of Epithelial Appendage Morphogenesis

51. Vainio S, Karavanova I, Jowett A et al. Identification of BMP-4 as a signal mediating secondary induction between epithelial and mesenchymal tissues during early tooth development. Cell 1993; 75:45-58. 52. Mitsiadis T, Lardelli M, Lendahl U et al. Expression of Notch 1, 2 and 3 is regulated by epithelial- mesenchymal interactions and retinoic acid in the developing mouse tooth and associated with determination of ameloblast cell fate. J Cell Biol 1995; 130:407-418. 53. Hogan BL. Bone morphogenetic proteins in development. Curr Opin Genet Dev 1996; 6:432-438. 54. Åberg T, Wozney J, Thesleff I. Expression patterns of bone morphogenetic proteins (BMPs) in the developing mouse tooth suggest roles in morphogenesis and cell differentiation. Dev Dyn 1997; 210:383-396. 55. Verschueren K, Dewulf N, Goumans MJ et al. Expression of type I and type IB receptors for activin in midgestation mouse embryos suggests distinct functions in organogenesis. Mech Dev 1995; 52:109-123. 56. Vaahtokari A, Vainio S, Thesleff I. Associations between transforming growth factor beta 1 RNA expression and epithelial-mesenchymal interactions during tooth morphogenesis. Development 1991; 113:985-994. 57. Chai Y, Mah A, Crohin C et al. Specific transforming growth factor-beta subtypes regulate embryonic mouse Meckel’s cartilage and tooth development. Dev Biol 1994; 162:85-103. 58. Wilkinson DG, Bhatt S, McMahon AP. Expression pattern of the FGF-related protooncogene int-2 suggests multiple roles in fetal development. Development 1989; 105:131-136. 59. Jernvall J, Kettunen P, Karavanova I et al. Evidence for the role of the enamel knot as a control center in mammalian tooth cusp formation: Non-dividing cells express growth stimulating FGF-4 gene. Int J Dev Biol 1994; 38:463-469. 60. Heikinheimo M, Lawshe A, Shackleford GM et al. FGF-8 expression in the post-gastrulation mouse suggests roles in the development of the face, limbs and central nervous system. Mech Dev 1994; 48:129-138. 61. Peters KG, Werner S, Chen G et al. Two FGF receptor genes are differentially expressed in epithelial and mesenchymal tissues during limb formation and organogenesis in the mouse. Development 1992; 114:233-243. 62. Kettunen P, Karavanova I, Thesleff I. Expression of different splicing variants of FGF receptors during early tooth development. Dev Genet 1998; in press. 63. Bitgood MJ, McMahon AP. Hedgehog and Bmp genes are coexpressed at many diverse sites of cell-cell interaction in the mouse embryo. Dev Biol 1995; 172:126-138. 64. Vaahtokari A, Åberg T, Jernvall J et al. The enamel knot as a signaling center in the developing mouse tooth. Mech Dev 1996; 54:39-43. 65. Iseki S, Araga A, Ohuchi H et al. Sonic hedgehog is expressed in epithelial cells during development of whisker, hair, and tooth. Biochem Biophys Res Commun 1996; 218:688-693. 66. Chen Y, Struhl G. Dual roles for patched in sequestering and transducing Hedgehog. Cell 1996; 87:553-563. 67. Partanen AM, Thesleff I. Localization and quantitation of 125I-epidermal growth factor binding in mouse embryonic tooth and other embryonic tissues at different developmental stages. Dev Biol 1987; 120:186-197. 68. Kronmiller, JE, Upholt WB, Kollar EJ. Expression of epidermal growth factor mRNA in the developing mouse mandibular process. Arch Oral Biol 1991; 36:405-410. 69. Miettinen PJ, Berger JE, Meneses J et al. Epithelial immaturity and multiorgan failure in mice lacking epidermal growth factor receptor. Nature 1995; 376:337-341. 70. Sibilia M, Wagner EF. Strain-dependent epithelial defects in mice lacking the EGF receptor. Science 1995; 269:234-238. 71. Mitsiadis TA, Henrique D, Ish-Horowicz D et al. Mouse Serrate-1 (Jagged-1): Expression in the developing tooth is regulated by epithelial-mesenchymal interactions and fibroblast growth factor-4. Development 1997; 124:1473-1483. 72. Artavanis-Tsakonas S, Matsuno K, Fortini ME. Notch signaling. Science 1995; 268:225-232. 73. Sonnenberg E, Weidner K, Birchmeier C. Expression of the met-receptor and its ligand, HGF-SF during mouse embryogenesis. In Hepatocyte Growth Factor-Scatter Factor

The Teeth as Models for Studies on the Development and Evolution of Organs

177

(HGF-SF) and the C-met Receptor. Goldberg ID, Rosen EM, eds. Basel: Birkhäuser Verlag, 1993:382-394. 74. Tabata MJ, Kim K, Liu JG et al. Hepatocyte growth factor is involved in the morphogenesis of tooth germ in murine molars. Development 1996; 122:1243-1251. 75. Weil M, Itin A, Keshet E. A role for mesenchyme-derived tachykinins in tooth and mammary gland morphogenesis. Development 1995; 121:2419-2428. 76. Luukko K, Moshnyakov M, Sainio K et al. Expression of neurotrophin receptors during rat tooth development is developmentally regulated, independent of innervation, and suggests functions in the regulation of morphogenesis and innervation. Dev Dyn 1996; 206:87-99. 77. Luukko K, Arumäe U, Karavanov A et al. Neurotrophin mRNA expression in the developing tooth suggest multiple roles in innervation and organogenesis. Dev Dyn 1997; 210:117-129. 78. Ahrens K. Die Entwicklung der menschlichen Zähne. Arb anat Inst Wiesbaden 1993; 48:169-266. 79. Butler PM. The ontogeny of molar pattern. Biol Rev 1956; 31:30-70. 80. Parker SB, Eichele G, Zhang P et al. p53-independent expression of p21Cip1 in muscle and other terminally differentiating cells. Science 1995; 267:1024-1027. 81. Jernvall J, Åberg T, Kettunen P et al. The life history of an embryonic signaling center: BMP-4 induces p21 and is associated with apoptosis in the mouse tooth enamel knot. Development 1998; 125:161-169. 82. Niswander L, Martin GR. FGF-4 expression during gastrulation, myogenesis, limb and tooth development in the mouse. Development 1992; 114:755-768. 83. Riddle RD, Johnson RL, Laufer E et al. Sonic hedgehog mediates the polarizing activity of the ZPA. Cell 1993; 75:1401-1416. 84. Francis PH, Richardson MK, Brickell PM et al. Bone morphogenetic proteins and a signalling pathway that controls patterning in the developing chick limb. Development 1994; 120:209-218. 85. Echelard Y, Epstein DJ, St-Jacques B et al. Sonic hedgehog, a member of a family of putative signaling molecules, is implicated in the regulation of CNS polarity. Cell 1993; 75:1417-1430. 86. Fan CM, Tessier-Lavigne M. Patterning of mammalian somites by surface ectoderm and notochord: evidence for sclerotome induction by a hedgehog homolog. Cell 1994; 79:1175-1186. 87. Liem KF Jr, Tremml G, Roelink H et al. Dorsal differentiation of neural plate cells induced by BMP-mediated signals from epidermal ectoderm. Cell 1995; 82: 969-979. 88. Nohno T, Ishikawa T, Saito T et al. Identification of a human type II receptor for bone morphogenetic protein-4 that forms differential heteromeric complexes with bone morphogenetic protein type I receptors. J Biol Chem 1995; 270:22522-22526. 89. Bellusci S, Furuta Y, Rush MG et al. Involvement of Sonic Hedgehog (Shh) in mouse embryonic lung growth and morphogenesis. Development 1997; 124:53-63. 90. Widelitz RB, Jiang TX, Noveen A et al. FGF induces new feather buds from developing avian skin. J Invest Dermatol 1996; 107:797-803. 91. Ting-Berreth SA, Chuong CM. Sonic hedgehog in feather morphogenesis—induction of mesenchymal condensation and association with cell death. Dev Dyn 1996; 207:157-170. 92. Vaahtokari A, Åberg T, Thesleff I. Apoptosis in the developing tooth—association with an embryonic signaling center and suppression by EGF and FGF-4. Development 1996; 122:121-129. 93. Coelho CN, Upholt WB, Kosher RA. The expression pattern of the chicken homeoboxcontaining gene GHox-7 in developing polydactylous limb buds suggests its involvement in apical ectodermal ridge-directed outgrowth of limb mesoderm and in programmed cell death. Differentiation 1993: 52:129-137. 94. Graham A, Francis-West P, Brickell P et al. The signalling molecule BMP4 mediates apoptosis in the rhombencephalic neural crest. Nature 1994; 372:684-686.

178

Molecular Basis of Epithelial Appendage Morphogenesis

95. Zou H, Niswander L. Requirement for BMP signaling in interdigital apoptosis and scale formation. Science 1996; 272:738-741. 96. Davidson EH. Later embryogenesis: regulatory circuitry in morphogenetic fields. Development 1993; 118:665-690. 97. Chen Y, Bei M, Woo I et al. Msx-1 controls inductive signaling in mammalian tooth morphogenesis. Development 1996; 122:3035-3044. 98. Satokata I, Maas R. Msx-1 deficient mice exhibit cleft palate and abnormalities of craniofacial and tooth development. Nat Genet 1994; 6:348-356. 99. Van Genderen C, Okamura RM, Farinas I et al. Development of several organs that require inductive epithelial-mesenchymal interactions is impaired in LEF-1-deficient mice. Genes Dev 1994; 8:2691-2703. 100. Zhou P, Byrne C, Jacobs J et al. Lymphoid enhancer factor 1 directs hair follicle patterning and epithelial cell fate. Genes Dev 1995; 9:700-713. 101. Kratochwil K, Dull M, Fariñas I et al. Lef1 expression is activated by BMP-4 and regulates inductive tissue interactions in tooth and hair development. Genes Dev 1996; 10:1382-1394. 102. Behrens J, Von Kries JP, Kuhl M et al. Functional interaction of beta-catenin with the transcription factor LEF-1. Nature 1996; 382:638-642. 103. Brunner E, Peter O, Schweizer L et al. Pangolin encodes a LEF-1 homologue that acts downstream of armadillo to transduce the wingless signal in Drosophila. Nature 1997; 385:829-833. 104. Peters H, Neubueser A. Balling R. Failure of tooth formation and craniofacial abnormalities in Pax9 deficient mice. Abstract in VIth International Conference on Tooth Morphogenesis and Differentiation, Göteborg, Sweden, June 11-15, 1997. 104a.Neubüser A, Peters H, Balling R, Martin GR. Antagonistic interactions between FGF and BMP signaling pathways: A mechanism for postioning the sites of tooth formation. Cell 1997; 90:247-255. 105. Matzuk MM, Kumar TR, Shou W et al. Transgenic models to study the roles of inhibins and activins in reproduction, oncogenesis, and development. Recent Prog Horm Res 1996; 51:123-154. 106. Morrison-Graham K, Schatteman GC, Bork T et al. A PDGF receptor mutation in the mouse (Patch) perturbs the development of a non-neuronal subset of neural crest-derived cells. Development 1992; 115:133-142. 107. Peterkova R, Peterka M, Ruch JV. Morphometric analysis of potential maxillary diastemal dental anlagen in three strains of mice. J Craniofac Genet Dev Biol 1993; 13:213-222. 108. Tureckova J, Sahlberg C, Åberg T et al. Comparison of expression of the Msx-1, msx-2, BMP-2 and BMP-4 genes in the mouse upper diastemal and molar tooth primordia. Int J Dev Biol 1995; 39:459-468. 109. Keränen SVE, Jernvall J, Thesleff I. Molecules associated with tooth loss in evolution— expression of developmental genes in vole diastema buds. Abstract in The Fifth International Congress of Vertebrate Morphology, Bristol, Britain, July 13-17, 1997. 110. Nieminen P, Arte S, Pirinen S. Gene defect in hypodontia—exclusion of msx1 and msx2 as candidate genes. Hum Genet 1995; 96:305-308. 111. Arte S, Nieminen P, Pirinen S et al. Gene defect in hypodontia: exclusion of EGF, EGFR, and FGF-3 as candidate genes. J Dent Res 1996; 75:1346-1352. 112. Vastardis H, Karimbux N, Guthua SW et al. A human MSX1 homeodomain missense mutation causes selective tooth agenesis. Nat Genet 1996; 13:417-421. 113. Kere J, Srivastava AK, Montonen O et al. X-linked anhydrotic (hypohydrotic) ectodermal dysplasia is caused by mutation in a novel transmembrane protein. Nat Genet 1996; 13:409-416. 114. Gruneberg H. Genes and genotypes affecting the teeth of the mouse. J Embryol exp Morph 1965; 14:137-159. 115. Gruneberg H. The glandular aspects of the tabby syndrome in the mouse. J Embryol exp Morph 1971; 25:1-19. 116. Sofaer JA. Aspects of the tabby-crinkled-downless syndrome. I. The development of tabby teeth. J Embryol Exp Morph 1969; 22:181-205.

The Teeth as Models for Studies on the Development and Evolution of Organs

179

116a. The Tabby phenotype is caused by mutation in a mouse homologue of the EDA gene that reveals novel mouse and human exons and encodes a protein (ectodysplasin-A) with collagenous domains. Proc Natl Acad Sci USA 1997; 94:13069-13074. 117. Blecher SR, Kapalanga J, Lalonde D. Induction of sweat glands by epidermal growth factor in murine X-linked anhydrotic ectodermal dysplasia. Nature 1990; 345:542-544. 118. Falconer DS, Fraser AS, King JWB. The genetics and development of ‘crinkled’, a new mutant in the house mouse. J Genet 1951; 50:324-344. 119. Philips RJS. Mouse News Letter 1960; 23:29. 120. Sofaer JA. The teeth of the “sleek” mouse. Arch Oral Biol 1977; 22:299-301. 121. Semina EV, Reiter R, Leysens NJ et al. Cloning and characterization of a novel bicoidrelated homeobox transcription factor gene, RIEG, involved in Rieger syndrome. Nat Genet 1996; 14:392-399.

CHAPTER 10

Epithelial-Mesenchymal Interactions in Gastrointestinal Development Drucilla J. Roberts

Introduction

T

he epithelial-mesenchymal interaction (EM) is a critical mechanism employed during embryogenesis. The signaling between these two different tissues is required to form the functional unit of the systems in which it is used. The gastrointestinal tract (gut) is dependent on EM interactions for its development and differentiation. The gut is a valuable model system to study EM interactions. The gut requires EM interactions from its initial organogenesis throughout the adult life of the organism. EM interactions are critical in the formation of gross pattern along the anterior-posterior (AP) axis, and local pattern along the crypt-villous (CV) axis. Many aspects of the development and pattern of the gastrointestinal tract are conserved among divergent animal species, suggesting a shared pathway of organogenesis. These similarities will be discussed herein and include: 1. initial developmental events in gut organogenesis; 2. the morphology of the gut; 3. molecular controls of these aspects of gut development and function. The gut is an early evolutionary innovation. The primitive tubular organism developed a tube within a tube, thus enjoying the remarkable ability to store and digest nutrients. The utility of an internalized digestive system relieved the organism of developmental constraints on body size and allowed for the ability to evolve other differentiated structures and systems. By forming an entry and exit to this system, the gut produced a patterned body plan with an anterior and posterior axis. Gut epithelium is derived from ectoderm and endoderm. Most of the gut is composed of definitive endoderm and splanchnic mesoderm in a tube-like arrangement. The epithelium in the most anterior region of the gut (for example the mouth) and most posterior region (for example the anus) is derived from ectoderm. The EM interactions discussed below will focus on the endodermally derived epithelium. The vertebrate gut is patterned along the anterior-posterior (AP) axis with morphologically and physiologically distinct regions. The gut as a system is critical not only in its function as a digestive organ, but embryologically the gut endoderm is an essential interactor required for the formation of the heart1-5 and for providing the anlage and signals to form many foregut derived organs. Gut derivatives are formed by EM interaction-directed budding morphogenesis. These include as derivatives the thyroid, lungs, liver and pancreas. The study of the gastrointestinal tract includes elegant descriptions of morphologic aspects of the gut during development in many species,6 classic experiments in which Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

182

Molecular Basis of Epithelial Appendage Morphogenesis

experimental embryology and cell culture techniques were used to explore the tissue specificity of the inductive signals critical in gut differentiation (see below), and recent descriptive and experimental work investigating the molecular controls of gut regionalization (described herein). This review will principally focus on the vertebrate gut, but one cannot discuss molecular controls of development without referencing the system to which we owe much of our genetic discoveries, Drosophila.

Initial Development of the Vertebrate Lumenal Gut: Inductive Events Organogenesis must proceed in a highly regulated temporal and spatial pattern to insure embryonic viability. The gut tube is formed with remarkable similarity in all vertebrates and many invertebrates.7 Critical in early gut formation is the invagination of the primitive endoderm and the subsequent growth and differentiation of the subjacent mesenchymal mesoderm. Tissue recombination studies described below have shown that the presumptive gut endoderm is capable of signaling the mesoderm, induce smooth muscle differentiation.8 Initially there is an anterior ventral invagination at the head fold stage. The ventral endoderm and subjacent mesenchymal mesoderm are internalized forming the anterior intestinal portal (AIP) (Fig. 10.1). The control of this event is poorly understood. AIP formation may be a passive response from head folding bringing the endoderm up ‘under’ the fold as a sequellae of growth of the brain,9 or may be an active endoderm specific function.9 Subsequently the splanchnic mesoderm closely associated with the endoderm undergoes smooth muscle differentiation as it forms a tube around the endoderm. This tube grows as an open cylinder, closing anteriorly as it elongates posteriorly. Shortly after the AIP is formed and has begun to elongate, the caudal intestinal portal (CIP) forms at the posterior end of the embryo (Fig. 10.1). The endoderm at the caudal end of the embryo invaginates inward and the subjacent mesenchymal mesoderm surrounds it, forming a tube similarly to the formation of the AIP. Although this invagination mirrors that of the earlier AIP invagination, it is even more poorly understood, as there is no simultaneously occurring body folding event. It has been suggested that the growth of the tail bud posteriorly may facilitate the CIP invagination.9 The growth and elongation toward each other result in the tubes meeting and fusing, closing around the connection to the yolk sac/ stalk in the middle of the embryo. Initially the gut tube is straight and uniform in its morphology, with no apparent gross or microscopic regional differences. Early in gut development an AP pattern is formed with regionally distinct sections: fore-, mid-, and hind-gut (Fig. 10.2). The first gross morphological regional distinction is the rotation and distention in the posterior foregut as the start of the differentiation of the stomach. The liver diverticulum and the lung buds begin their development from the foregut at about this stage as well. Following this is elongation and looping of the midgut and (usually) the formation of short tubal structures at the midgut/ hindgut boundary (appendix in mammals, ceca in birds). These events occur before the ventral body wall closes, at limb bud stages (by stage 23 in the chick).10,11 Therefore, in very early developmental stages the three regions of the gut are formed: foregut from the AIP, hindgut from the CIP, and midgut from both AIP and CIP (Fig. 10.2). The mature gut pattern forms later with continued growth, characteristic rotations, and organ specific cellular differentiation in both the endoderm and mesoderm components. It has been known for decades that the gut cannot develop normally without an interaction between the endoderm and mesoderm.12-15 The inductive interaction has been shown to be reciprocal. When isolated endodermal tissue is dissected from the primitive gut tube and cultured in isolation, tissue growth is achieved without differentiation. But when endoderm is cultured associated with its mesoderm, morphologic development is possible, and

Epithelial-Mesenchymal Interactions in Gastrointestinal Development

183

Neural folds beginning to fuse Fore gut

Heart

Anmion (cut) t Midgut Hindgu

Pericardial coelom Anterior intestinal port

Yolk sac

Posterior intestinal portal

Body stalk

Fig. 10.1. AIP/CIP. Diagram of a midsagittally sectioned chick embryo at approximately stage 1410 showing the initial stages of gut development via invagination forming the anterior intestinal portal and posterior intestinal portal. Anterior is to the left, ventral is down.

foregut

kidney

midgut

Fig. 10.2. Vertebrate gut anatomy. Drawings of approximately stage 19-22 (top) and stage 26-28 (bottom) whole chick embryos with ventral wall open and membranes removed. (Adapted from Romanoff11).

yolk stalk hindgut

lung bud proventricularis yolk stalk

gizzard kidney

cloaca

ceca

small intestine

gut specific morphology is seen.13-15 The direction of these inductive events has been the focus of much work. While primitive foregut endoderm can differentiate to form gastric glands when cultured with primitive foregut mesoderm, this result can also be obtained when the same endoderm is cultured with skin fibroblasts.13,16 This might indicate that the endoderm is competent and the mesoderm provides nonspecific support of differentiation. Alternatively, the endoderm may induce visceral muscle differentiation, e.g., to form smooth muscle which subsequently can induce or permit epithelial differentiation. As it turns out, both are partially true. There is a developmental time window after which the primitive gut

184

Molecular Basis of Epithelial Appendage Morphogenesis

endoderm, although still morphologically undifferentiated, is committed and can develop into its regionally specific epithelium when cultured with a variety of tissues, including the vitteline membrane.17 But the same primitive endoderm cultured with nongut derived mesoderm (e.g., skin fibroblasts), in turn stimulates smooth muscle development as assayed by histology and by induction of visceral mesodermal proteins, e.g., tenascin18 and smooth muscle actin.16 Whether this transformed mesodermal tissue can subsequently induce precommitment stage primitive endoderm is unknown.

Patterning and Epithelial Differentiation: Inductive Events There is a general correlation of the epithelial pattern (cellular phenotype) and gross organ morphology with the AP region of the body plan. At the gross level, the AP regional differences of the gut are clearly apparent. Differences in the diameter of the tube, position, rotation, orientation of the smooth muscle surrounding the tube, as well as location of gut derivatives, are all key landmarks in the AP pattern. A common AP gut pattern exists such that for most species: esophagus and stomach derive from the foregut, small intestines derive from the midgut, and colon derives from the hindgut. The controls of the AP regionalization of gut morphogenesis are unknown. As these regions generally relate to the overall body plan (foregut in thorax and upper abdomen, hindgut in abdomen and pelvis), the genetics involved in body plan and segmentation are likely important in regional gut specification as well. The cytodifferentiation of the epithelium also correlates to the AP pattern, with specific microscopic and cellular protein phenotypes per each region (Fig. 10.2). As the mesoderm becomes committed to its AP regional fate, it can influence the ultimate epithelial AP pattern. When primitive gut endoderm is dissected from the early gut tube, separated by region (e.g., into fore-, mid-, and hindgut endoderm), and recombined as explants with gut derived mesoderm (also separated by region), the endoderm will adopt an epithelial fate of the mesodermal region with which it has been combined.8,13,14,19 For example, stomach endoderm cultured with stomach mesoderm differentiates with a stomach epithelial phenotype (homologous differentiation). Yet when stomach endoderm is cultured with small intestinal mesoderm a small intestinal epithelial phenotype results (heterologous differentiation). Therefore, the mesoderm can direct the phenotype of the endoderm (Fig. 10.3). The time window of endodermal competency for this response is limited. The endoderm becomes committed to its AP epithelial phenotypic fate long before regional epithelial morphologic differences are evident. In the chick, gut epithelial differentiation begins at about day 5-6 of incubation (of a 21 day gestation11) but commitment occurs probably by day 2.5.17,20 Chick endoderm retains some plasticity for heterologous differentiation up to day 6.13,21-24 In the rat this commitment occurs by day 16.5,16,25 and murine endodermal commitment occurs before 11.5 days.26 Although it has been generally accepted that the mesoderm directs endodermal morphologic differentiation, the epithelial morphological differentiation does not always equate with cytodifferentiation. The chick gut includes a two part stomach with the upper stomach (proventricularis) functioning in acid digestion and the lower stomach (gizzard) as a grinding/muscular organ. Each region of the two part stomach is distinct in its morphology (epithelial and mesodermal) and epithelial cytodifferentiation (cellular protein expression). The gizzard epithelium is aglandular whereas the proventricular epithelium is glandular. When chick presumptive gizzard endoderm is cultured with proventricular mesoderm, the gizzard endoderm develops glands and the epithelium will express a proventricular specific enzyme: pepsinogen.23,24,27 As predicted, in the reverse experiment in which presumptive proventricular endoderm is cultured with gizzard mesoderm, the epithelial morphology is gizzard and the cells do not express pepsinogen.24,27 But the mesodermal inductive capability does not always hold true for both morphological and cytological differentiation. Small

Epithelial-Mesenchymal Interactions in Gastrointestinal Development

185

Fig. 10.3. Technique of culture. Diagram of dissociation of the endoderm from the mesoderm of an approximately stage 13 chick embryo ventral view. Mechanical dissociation following proteinase digestion is followed by culture of homologous or heterologous (shown) regions in a humidified chamber. (Adapted from refs. 12, 21).

intestinal epithelium has a microscopic morphology distinct from proventricular. The epithelium is composed of a mucosa and submucosa forming villous projections into the lumen of the small intestine. The small intestinal epithelium expresses enzymes distinct from the proventricularis. When presumptive small intestinal epithelium is cultured with proventricular mesoderm, the morphology is predictably glandular (proventricular), but the cells do not express pepsinogen.23,28,29 This is true even when very early (15-20 somite stage, E2, approximately stage 13) endoderm is used.30 The converse experiment has the expected results of proventricular endoderm differentiating toward a small intestinal fate, including formation of villous structures and expression of sucrase (a small intestinal marker).30 A summary of this data is provided in Table 10.1. It is clear that nonproventricular endoderm can differentiate to proventricular morphology and can produce proventricular specific proteins when under the influence of proventricular mesoderm. The fact that gizzard epithelium can be influenced in this manner when it is not in normal development suggests two possible explanations. One is that an inductive positive signal is present in the proventricular mesenchyme that influences both glandular development and cytodifferentiation in the overlying endoderm. This would explain the ability of the proventricular mesoderm to induce gastric glands in the otherwise aglandular gizzard epithelium. There would have to be absence of this signal in the gizzard mesoderm, as the proventricular endoderm does not form glands when apposed to this mesoderm. Others have suggested that the gizzard mesodermal influence is inhibitory.27,31

Molecular Basis of Epithelial Appendage Morphogenesis

186

Table 10.1. Heterologous organ explant experiment Endoderm

Mesenchyme

Mesenchyme Morphology

Gizzard Proventricularis Proventricularis Small intestine

proventricularis gizzard small intestine proventricularis

glands squamous villi glands

Pepsinogen+

Sucrase+

+ – – –

– – + +/–

Adapted from Yasugi et al, 1991.15

The mesenchymal influence on endoderm patterning primarily involves specification of morphology, which may not include cytodifferentiation. The difference between midgut and fore/hindgut endoderm’s ability to undergo heterologous differentiation may be an endogenous characteristic, as it is evident in chicks (as described above) and rodents.32 It may be that, unlike foregut and hindgut endoderm, the midgut endoderm is self-determined at least as far as cytodifferentiation. The mesoderm thus signals the endoderm and affects (at least) epithelial morphologic phenotype. What is this signal? Direct EM cell-cell contact has been noted.33 In some species cell-cell contact is required for gut induction (for example C. elegans34 and Drosophila35). In vertebrates, transfilter experiments in which cell to cell contact was inhibited showed heterotypic differentiation, suggesting that cell-cell contact is not essential and that secreted factors must play a critical role.18,25,36 Other lines of research have focused on the role of the extracellular matrix proteins in directing gut differentiation via EM.15,37,38 A basement membrane is present at the interface of the epithelium and mesoderm throughout the length of the gut. In fact, the interaction of the endoderm and mesoderm is required for the formation of the basement membrane which separates them. Either cell type cultured alone fails to produce a basement membrane, and in either homotypic or heterotypic coculture experiments the basement membrane has to be reconstituted before epithelial cytodifferentiation begins.37,39-41 Components of the basement membrane are deposited by both the epithelium and mesoderm as detected by cross-species co-cultures using species specific antibodies.42,43 Here we see a common problem in the understanding of how differentiation can be directed by molecules; there is a difference between local and global patterning. Although a gradient of extracellular matrix proteins exists along the CV axis (an example of local patterning),42 the basement membrane/extracellular matrix proteins appear to be uniform in their constituents and distribution throughout the AP axis of the gut (an example of global patterning). Therefore local patterning may be influenced via the basement membrane/extracellular matrix proteins, but this is unlikely to play a direct role in global pattern.44-46 The importance of this tissue which separates and joins the gut epithelium and mesoderm cannot be ignored in directing the AP pattern, as presumably inductive molecules are transmitted via these proteins.

Genetic Controls of Gut Development Compared to the decades of work using coculture techniques in the study of gut development, molecular techniques have only recently been applied to this area.

Epithelial-Mesenchymal Interactions in Gastrointestinal Development

187

Despite the enormous advances in understanding the genetic controls of patterning the basic body plan, relatively little work has focused on visceral patterning. In the gut, this may be a function of the methods employed to analyze gene expression and phenotypes of genetically mutant vertebrate embryos. Most murine developmental geneticists examine phenotypes by sagittal or coronal histologic sections, or by clearing and staining embryos to examine the skeleton. Obviously these techniques are suboptimal for evaluation of the lumenal gastrointestinal tract, as the AP orientation and boundaries are nearly lost in sections and are destroyed in skeletal preparations. Much more is known about the molecular regulation of gut development in Drosophila.

Genetics of Gut Development in Drosophila The fly gut is remarkably similar to vertebrate gut in its morphology (AP regions), formation via invaginations, dependency on EM interactions, and as we will see, in some of the genetic controls of its development. The fly gut AP regions include a fore-, mid-, and hind-gut (Fig. 10.4). The foregut is composed of a pharynx, esophagus, and proventricularis (stomach). The midgut includes the gastric ceca and intestine. The hindgut also receives the exit of the Malpighian tubules (similar to vertebrate kidneys). All three regions develop from separate invaginations which grow together to form an intact tube. Whereas Drosophila fore- and hind-gut epithelium is ectodermally derived, the midgut is endodermally derived. The midgut in Drosophila is demarcated anteriorly by the gastric ceca and posteriorly by the Malpighian tubules (bordering the hindgut) (Fig. 10.4). Drosophila’s midgut is characterized by three mesodermal constrictions, which are phenotypic markers for genetic control. Analysis of midgut development has shown that, as is true in vertebrates, the midgut epithelial development is dependent on EM interactions. Examination of flies with mutations resulting in absent or aberrant development of the visceral mesoderm fail to develop midgut epithelium (as in twist, and twist snail double mutants35,47,48). It appears that the endoderm must have direct contact with the mesoderm to form epithelium. This interaction likely involves the cadherin family of membrane proteins, one of which has been identified—shotgun,35 which encodes Drosophila E-cadherin.49 Shotgut mutants have malformed midguts.35,49 An important class of embryonic control genes involved in Drosophila midgut development are the Homeotic (Hom) genes, encoding homeobox containing transcription factors, key regulators of the embryonic body plan.50-52 Hom genes are classified for their function in designating body regions. Hom mutants have alterations in the basic body plan, some causing ‘homeosis’ of one region to another (for example in ultrabithorax the abdomen is homeotically transformed into a second thorax50). Genetic dissection of Drosophila midgut morphogenesis has delineated a pathway in which the restricted expression of specific Hom genes controls pattern in the Drosophila midgut by demarcating and regulating the formation of the midgut constrictions53-57(Fig. 10.5). While most of the Hom genes with patterning roles in Drosophila midgut are mesodermally restricted in expression (Scr, Antp, Ubx, Abd-A, Abd-B), Labial (lab) is expressed in a restricted region of the midgut endoderm, critical for the second constriction and endodermal cell fate (required for the formation of copper cells58). Downstream targets of the Hom genes controlling midgut patterning include teashirt (tsh), also a Hom gene required for proper formation of the anterior and central midgut structures.59 Tsh, mesodermally expressed, is a target of three Hom genes (Ubx, abd-A, and Antp). Tsh is also regulated by two mesodermally expressed secreted factors: decapentaplegic (dpp), which encodes a TGF-β superfamily protein, and wingless (wg).53,57 These secreted

188 Fig. 10.4. Drosophila larval gut anatomy. Ventral view; anterior is up. (Adapted from Bate and Arias61).

Molecular Basis of Epithelial Appendage Morphogenesis

pharnix

salivary duct imaginal ring salivary gland

esophagus foregut imaginal ring gastric caecum

proventriculus midgut midgut imaginal histoblasts

hindgut imaginal ring

Malpighian tubule hindgut

anal imaginal ring

Fig. 10.5. Genetic controls of Drosophila midgut morphogenesis. (Adapted from Roberts et al74).

Epithelial-Mesenchymal Interactions in Gastrointestinal Development

189

proteins signal between the visceral mesoderm and the endoderm, a key event in formation of the midgut constrictions. One method in which these secreted factors have been shown to signal the endoderm is by altering the subcellular localization of yet another Hom gene, extradenticle (exd).60 Drosophila fore- and hindgut development is fundamentally different than midgut development; they are ectodermally derived structures with less well documented EM tissue interactions. Although the genetic basis for regional specification in the fore- and hind-gut is poorly understood, early events in regionalization are known to be controlled by maternal effect genes (expression of which in the egg from the ovary sets up morphogenic gradients) which establish the primordia and axes of the gut.61 Zygotic control of fore- and hindgut development involves activation of the ectodermally expressed gene forkhead (fkh), a member of a class of transcription factors which includes many vertebrate homologues (see below). Fkh acts homeotically in the specification of terminal gut ectodermal tissues and plays a role in gut development.61 Once the fore- and hindgut primordia are specified, subsequent regionalization is accompanied by activation of patterning genes. In the hindgut, patterning involves activation of caudal (cad), a Hom gene with a pivotal role in regionalization of posterior embryonic elements.62 In the foregut, the genetics of patterning has recently been described for the proventricularis. Genes involved in proventricular morphogenesis include cell surface adhesion molecules, a zinc-finger transcription factor encoded by cubitus interruptis (ci), and secreted factors encoded by hedgehog (hh), wg, and dpp. It is of particular interest that signaling in Drosophila foregut patterning involves an interaction in which hh, expressed in the epithelium, is upstream of dpp, expressed in the mesoderm, and required for patterning the proventricularis which demarcates the foregut/midgut boundary in the foregut.63 In hh mutants the normal expression of dpp in the midgut structures at the foregut/midgut boundary (gastric ceca) is lost.63 As many genes known to play important roles in Drosophila development have homologues in vertebrates with similar functions, it is reasonable to assume that homologues for the genes discussed above play a role in the EM interactions involved in vertebrate gut development.

Genetic Controls of Vertebrate Gut Pattern Since the AP pattern of the vertebrate gut has a positional relationship to the overall pattern of the AP body plan, one might ask if the Hox genes, vertebrate homologues of Drosophila Hom genes, are involved in regulating gut AP pattern. Hox genes probably emerged via quadruplication of an ancestral homeobox gene cluster in the vertebrate lineage. Four homologous complexes, HoxA, HoxB, HoxC, and HoxD exist each on different chromosomes. Homeodomain sequence homologies have led to classification of Hox genes into subfamilies, each containing 2-4 related paralogous genes64 (Fig. 10.6). Genes within a cluster are expressed with spatial and temporal colinearity. The 3' genes are expressed earlier and in more anterior domains, and the 5' genes are expressed later and in more posterior domains.65 Hox genes have been shown to be key regulators of AP pattern in the limb bud, axial tissues, and the hindbrain.66,67 In each of these tissues the Hox genes are expressed in overlapping anteroposterior domains which correlate with structural boundaries. The analysis of Hox genes and their roles in patterning these regions have focused primarily on their role in skeletal pattern and pattern formation of the hindbrain. The analysis of Hox gene expression or function in visceral pattern is a recent area of investigation. Visceral homeosis has been described in the uterus of murine HoxA10 mutants68 and the vas deferens of HoxA11 mutants.69 Visceral anomalies have been described in the athymic and

190

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 10.6. Vertebrate Hox genes aligned chromosomally with their Drosophila homologs.

Fig. 10.7 (opposite). Hox hindgut boundaries. (Top) Diagrammatic representation of vertebrate gut morphogenesis with the anterior intestinal portal and caudal intestinal portal growing and elongating (large arrows) towards the umbilicus (dashed horizontal line). The regionalization of the lumenal gut forms foregut, midgut, and hindgut derived from the invaginations of the AIP (foregut and midgut) and CIP (midgut and hindgut). Cytodifferentiation of these regions forms the small intestine from the midgut, the ceca from both midgut (anteriorly) and hindgut (posteriorly), and large intestine and part of the cloaca from hindgut. The regionally restricted pattern of expression of the Abd-B-like Hox genes demarcates morphologic distinctions in the midgut and hindgut visceral mesoderm, diagrammatically shown with anterior limits of expression noted (exception is HoxC9, posterior limit of expression shown). (Bottom) mRNA expression patterns of the Abd-B-like Hox genes are shown by whole mount in situ hybridization. Paralogues are aligned. There is no paralogue of HoxD12 in the HoxA cluster. Paralogues without detectable hindgut expression (HoxB9, HoxC10, HoxC11) are not shown. HoxC12 and HoxC13 were not studied. Expression limits in the visceral mesoderm can be seen around the midgut and hindgut boundary of the ceca and the posterior limit of the hindgut, the cloaca. The ceca in each panel are highlighted by the thin white lines. In general, paralogues of different clusters exhibit similar anterior expression borders in the hindgut as they do in other embryonic tissues. HoxD9 in the gut is an exception to this rule, as it is expressed in a domain seemingly identical to HoxA10 and HoxD10 at this stage. Expression of HoxA13 and HoxD13 is also found in the endoderm (long arrow indicates endodermal expression, arrowhead notes mesodermal expression. The anterior extent of HoxD13' s endodermal expression is not evident in this photograph). AIP = anterior intestinal portal, Ce = ceca, CIP = caudal intestinal portal, Cl = cloaca, EN = endoderm, FG = foregut, MG = midgut, HG = hindgut, LI = large intestine, SI = small intestine, VM = visceral mesoderm. (Adapted from: Roberts et al74).

Epithelial-Mesenchymal Interactions in Gastrointestinal Development

A9

A10

A11

191

A13

C9

D9

Fig. 10.7.

D10

D11

D12

D13

192

Molecular Basis of Epithelial Appendage Morphogenesis

hypothyroid phenotype of HoxA3 mutants.70 As discussed below, there is expression and experimental data supporting a role for the Hox genes in gut patterning and development. Several Hox genes have been noted to be expressed in the vertebrate gut, including the HoxD genes in the Abd-B class.71-73 5' HoxD gene expression has been described in the mesoderm of the posterior midgut and hindgut. Due to the sectioning method of in situ analysis, precise AP boundaries of expression could not be assessed. Recent work describing Hox gene expression in the chick gut looked at expression in early embryonic stages by whole mount in situ hybridization.74,75 In the chick hindgut the Abd-B-like Hox genes of the A and D cluster have regionally restricted expression boundaries demarcating morphologic landmarks of the mid- and hind-gut74,75 (Fig. 10.7). In general, the genes are expressed in a ‘nested’ pattern, with sharp expression limits anteriorly and overlapping expression at the most posterior hindgut region (Fig. 10.7), reminiscent of the expression pattern of the Abd-B like HoxD genes in the posterior region of the limb bud.76,77 The embryonic day 4 (E4) expression patterns suggest that the restricted boundaries of expression of the Abd-B-like Hox genes demarcate the regions which will form the cloaca, large intestine, ceca, mid-ceca at the midgut/hindgut border, and the posterior portion of the midgut (Fig. 10.7). Moreover, these molecular events presage regional distinctions. Expression of all Hox genes could be detected by stage 14, well before the hindgut lumen is closed (by stage 28). Cytodifferentiation of the hindgut mesoderm and epithelium begins later, at stages 29-31.11

The Expression Patterns of Hox Genes in the Hindgut Are Dynamic The expression domains of HoxA10 and HoxD10, which originally extend to the posterior limit of the gut, resolve into restricted domains solely encompassing the ceca.74 Once the nested expression domains of the Abd-B related Hox genes are established, their relative anterior limits of expression are maintained through subsequent growth and differentiation. The refinement of the Hox expression patterns may involve cross-regulation between the Hox genes, or may be due to secondary factors. Although all the genes analyzed were expressed in the mesoderm, with the 3' paralogues expressed more anteriorly than the 5', the 13th paralogue was unique in its additional endodermal expression. Both HoxA13 and HoxD13 are expressed in the mesoderm of the chick cloaca (the common gut and urogenital orifice); they are also both expressed in the endoderm of the hindgut.74 At E4-5 there is a sharp anterior endodermal expression boundary at the cecal buds. As Hox genes are known to regulate pattern in other regions of the developing embryo via restricted expression pattern, and restricted expression in the gut is observed, it is reasonable to assume that Hox genes play a role in regulating the AP pattern of the gut. What functional evidence do we have that supports this suggestive expression pattern? A role for Hox genes in gut morphogenesis has been implicated by analysis of Hox gene misexpression and mutation studies. These reports suggest that Hox genes might be involved in controlling vertebrate gut pattern. In an early study, a transgenic HoxA4 mouse mutant was developed in which a portion of the endogenous 3' untranslated region was substituted with simian virus 40 (SV40) T-antigen coding region and poly(A) addition site such that overexpression of endogenous HoxA4 resulted.78 The phenotype described was one of obstipation with colonic dilatation. HoxA4 is not normally expressed in the colon, but expression has been described in the mesoderm of the midgut.79-81 The effect on the hindgut must be a secondary effect of overexpression in the midgut (and perhaps ectopic expression in the hindgut). HoxC4 is expressed in the esophagus,82 although its regional expression pattern in the gut has not been well described. A targeted disruption of HoxC4 was found to cause an

Epithelial-Mesenchymal Interactions in Gastrointestinal Development

193

esophageal defect.83 In the HoxC4 -/- mouse the esophagus was not functional, and the authors ascribed the defect to esophageal obstruction due to malformation of the muscular layers of the distal esophagus and overproliferation of the epithelium. The data presented in the paper shows a poorly organized external muscular layer and a collapsed lumen. Loss of HoxC4 was also associated with changes in the expression of HoxC5 and HoxC6. Pollock et al describe the phenotype of a transgenic mouse in which HoxC8 was anteriorly misexpressed under the HoxA4 promoter.84 The mice had vertebral homeotic malformations and, additionally, specific gut maldevelopment was noted. The murine posterior stomach epithelium had differentiated as duodenal (anterior midgut) epithelium (a homeotic malformation). This finding is important, as it suggests that misexpression of a Hox gene in the mesoderm results in a transformation of the epithelium (EM interaction). There is a demonstration that murine HoxD12 and HoxD13 play a role in hindgut development. As we found in the chick,74 these two Hox genes are expressed in the most posterior region of the hindgut in the mouse, with overlapping mesodermal expression in the anal sphincter muscle (the posterior-most mesodermal level of the rectum). Kondo et al describe the gut phenotype of HoxD12 and HoxD13 mutant homozygotes.85 Homozygous mutants for HoxD12 showed a subtle rectal phenotype demonstrable only by histologic section of the region. Microscopically the anal sphincter muscle was disorganized and thin. The same finding was even more pronounced in the HoxD13 mutant homozygotes and often resulted in anal/rectal prolapse due to dysfunction of the sphincter. Despite all the work done over the last 10 years or so of analyzing Hox gene function by mutation and misexpression, only those described above have noted gut effects. It is likely an artefact of the type of analysis performed to examine the phenotype (e.g., by sectioning). But, this may support the premise that Hox genes function by patterning the overall body plan (global signaling) but not specific organs or systems (local signaling). Another interpretation is that as four complexes and 13 paralogous groups exist, a redundancy of function among Hox genes could protect against visceral malformations in single Hox gene ‘knockout’ experiments. This latter interpretation is supported by the compound mutant homozygotes. Unfortunately most of the compound Hox mutant studies have focused on the skeletal malformations and have not described any visceral findings (for example, refs. 86, 87). Despite this, there are examples in which visceral anomalies are described supporting the role of Hox genes in visceral patterning. Renal morphogenesis, although not a gut derivative, also requires EM interactions for normal development.88,89 In both the HoxA11 and HoxD11 single mutant homozygotes, normal kidneys are formed,90 yet the compound mutant homozygotes have severe renal hypoplasia.91 The conservation of Hox gene expression in the gut of chick and mouse implies a conservation of function. In fact, the evolutionary conservation of expression throughout the gut is such that the most 5' of the Abd-B like Hox genes are expressed in the most posterior region of the gut of many species.85,92,93 The most primitive or ancestral function of the Hox genes may be to set up the first AP polarity of the organism. The most posterior of the Hox genes are expressed in the most posterior part of the hindgut in various metazoans studied.85 One can argue that the digestive system is the most primitive polar organ system and that the genetics of the AP polarity in this system were coopted for use in subsequently evolved systems, neural and limb.85 Kondo argues this point using the temporal colinearity attribution of the Hox genes.85 They suggest that the earliest expression pattern indicates its most ancestral function.85,93 It is intriguing as well that organisms without polarity to their gut appear not to have a Hox gene or genes of the Abd-B class.85,94,95 Although the discovery of downstream targets of Hox genes have proved elusive, an activator has been identified. Sonic hedgehog, a vertebrate homologue of Drosophila’s hh, has been shown to be an upstream activator of HoxD genes in the vertebrate limb.96 Sonic is

194

Molecular Basis of Epithelial Appendage Morphogenesis

a signaling molecule implicated in mediating pattern in several regions of the embryo including the limb bud,96 somite,97,98 and neural tube.99-101 There is evidence that Sonic can act as a signal in EM interactions. In the limb bud, Sonic is expressed in the posterior medial mesoderm and is required for maintenance of the ectodermal expression of FGF-4.74,102,103 The protein product of the Sonic hedgehog gene, a secreted factor, is an excellent candidate for an early endodermally derived inductive signal in gut morphogenesis. Sonic (or its close relative Indian hedgehog) is expressed in endodermal tissues of the gut and its derivatives.104,106 The earliest endodermal expression of Sonic is restricted to the endodermal lips of the AIP and CIP.74 At this stage in the chick (approximately stage 13) the Abd-B like Hox genes are expressed in an nested mesodermal cluster around the Sonic-expressing endoderm,74 in a manner reminiscent of Sonic in the ZPA of the developing limb bud.96 In fact, just as virally mediated Sonic misexpression in the limb bud induces ectopic expression of the Abd-B like Hox genes,96,102 misexpression in the stage 10 presumptive midgut region of the embryo induces expression of HoxD11 and HoxD13.74 This induction appears to be sensitive to the AP level of the injection site, as ectopic expression of HoxD11 or HoxD13 was not detected when Sonic virus is injected anterior to the vitelline veins. If Sonic expression is required for gut invagination or Hox gene induction in the vertebrate gut, one would predict that if Sonic null mutants were viable up to gut developmental stages, a gut would fail to form. Mice with functionally null Sonic gene function are viable and apparently do form an intact gut (see ref. 105, Figures 2d and 5l). Could this be an example of redundant function, with the normally active Indian hedgehog gene functioning in the hindgut tissues? This has yet to be explored. Although the Sonic mutant mice form a gut, foregut abnormalities were noted. The esophagus is malformed with an enlarged lumen and disorganized or absent subjacent mesoderm (C. Chiang, personal communication). As we are proposing that Sonic is an endodermal signal important in EM interactions, this is an exciting supportive finding. In each organ in which the endoderm derived tissue expresses Sonic or Indian, there is closely associated mesenchymal mesoderm which expresses a homologue of dpp.74,104,106 There are two vertebrate homologues of dpp, Bmp-2 and Bmp-4. In the primitive hindgut, at the earliest time point in which Sonic expression can be detected in the CIP region (even before invagination is apparent), Bmp-4 expression is detectable in the subjacent mesenchymal mesoderm (Fig. 10.8).74 In Drosophila, at the foregut/midgut boundary hh is upstream of dpp.63 When Sonic is misexpressed using a retroviral vector into the midregion of the chick, ectopic Bmp-4 expression is induced in the mesoderm.74 Bmp-4 is also expressed adjacent to Sonic in the developing lung74,104,106 and plays a role in patterning the lung during development.107 Although Bmp-4 expression is not described in the Sonic knock out,105 lung abnormalities were noted. Targeted mutagenesis of Bmp-4 in the mouse resulted in early embryonic lethality due to failure of mesodermal development.108 In these embryos the ventral body wall failed to close and gut development was not observed. It may be that Bmp-4 is required for visceral mesodermal differentiation. There are intriguing parallels emerging between the morphologic development of the vertebrate and Drosophila gut and the molecules that play a role in their development. This suggests that since the last common ancestor between arthropods and chordates, aspects of the EM interactions and signaling cascade controlling gut development have been conserved. The conservation of the hh-dpp/Sonic-Bmp-4 pathway, as EM interacting signals seen in Drosophila and in the vertebrate gut, indicates that it is an ancient pathway. The convergence of this pathway and the induction of expression of Abd-B like Hox genes by Sonic in vertebrates has helped elucidate the downstream targets of these signals.

Epithelial-Mesenchymal Interactions in Gastrointestinal Development

A

195

B

Fig. 10.8. Bmp-4/sonic expression pattern. Sonic expression is detected in the endoderm of the AIP and CIP in pregut closure stage embryos seen at low magnification (left panel), ventral view. Endodermal expression confirmed in section (see Roberts et al74). At later stages Sonic is expressed in the gut in all levels (fore-, mid-, and hindgut) restricted to the endoderm. Bmp-4 is expressed in the mesoderm adjacent to Sonic at the CIP shown at low magnification (right panel), dorsal view. Mesenchymal expression is confirmed in section (see Roberts et al74). Bmp-4 gut expression persists but weakens in later stage embryos in the visceral mesoderm only.

Candidate upstream activators of Sonic include vertebrate homologues of the fkh gene class, identified in Drosophila. fkh encodes a novel transcription factor required for foreand hind- gut development. There are at least six vertebrate fkh homologues,109 all sharing homology with the DNA binding domain termed the forkhead domain. Three were originally identified for their importance in hepatic development: HNF-3α, β, and γ. All three are expressed in the murine gut endoderm with slightly different expression patterns along the AP axis of the gut.110,111 Detailed spatial/temporal expression patterns of the three have not been published, but HNF-3β is expressed in Hensen’s node at the site of the origin of the definitive endoderm110 and may be responsible its induction.112 In the mouse, HNF-3β is expressed at both the AIP and CIP.110,112,113 It has been suggested that HNF-3β regulates the production of Sonic in the notochord and floorplate99,100,110 and interacts with Sonic at the node.114 Chiang et al105 found that a reciprocal interaction exists in the maintenance of both Sonic and HNF-3β expression in the neural tube but also found an early Sonic-independent HNF-3β expression in late streak to early head fold stages. Late HNF-3β expression was found to be dependent on Sonic expression. HNF-3β may similarly lead to transcription of Sonic within the AIP/CIP endoderm. In preliminary studies we found HNF-3β expression limited to the chick AIP before expression of Sonic (data not shown). We find no expression at the CIP in stages studied. This difference in AP expression is consistent with a role for HNF-3β as an activator of Sonic at the AIP, or alternatively may play a role in modulating the AP regionalization of Sonic’s signal (such that although Sonic is expressed in both the AIP and CIP, the AIP alone has both Sonic and HNF-3β, differentiating the fore- from the hind-gut regions). Perhaps one of the other HNF-3 genes plays a similar regulatory role at the chick CIP. Other candidates for the genetic control of EM interactions in the gut include Nkx-2.3 and 2.5,115 Barx1,116 DCC,117 endomorphin,118 HGF/SF and c-Met (see for example ref. 119),

196

Molecular Basis of Epithelial Appendage Morphogenesis

Pax9,120 and members of the vertebrate homologues of the Cdx family,121-130 and Pdx-1.131-133 All of these genes have expression patterns or null mutant phenotypes which strongly suggest a role in gut regional development or differentiation. Gut induction and development has proved to be a useful model system to study EM interactions. As the molecular pathway is being elucidated, the focus of future study into the different molecular controls of global vs. local signaling in EM interactions should reveal new insights into the genetic controls of pattern regulation in global (overall body plan) and local (microscopic development) environments. We expect this field to be active in the coming years.

References 1. Jacobson AG, Sater AK. Features of embryonic induction. Development 1988; 104:341-359. 2. Sugi Y, Lough J. Anterior endoderm is a spcific effector of terminal cardiac myocyte differentiation of cells from the embryonic heart forming region. Dev Dyn 1994; 200:155-162. 3. Lough J, Barron M, Brogley M et al. Combined BMP-2 and FGF-4, but neither factor alone, induces cardiogenesis in non-precardiac embryonic mesoderm. Dev Biol 1996; 178:198-202. 4. Nascone N, Mercola M. An inductive role for the endoderm in Xenopus cardiogenesis. Development 1995; 121:515-523. 5. Sugi Y, Markwald RR. Formation and early morphogenesis of endocardial endothelial precursor cells and the role of endoderm. Dev Biol 1996; 175:66-83. 6. Hilton WA. The morphology and development of intestinal folds and villi in vertebrates. Am J Anat 1902; 1:459-504. 7. Gilbert SF, Developmental Biology. Fourth ed, ed. AD Sinaeur. 1994, Sunderland, Massachusetts: Sinauer Associates, Inc. 894. 8. Kedinger M, Simon-Assman PM, Lacroix B et al. Fetal gut mesenchyme induces differentiation of cultured intestinal endodermal and crypt cells. Dev Biol 1986; 113:474-483. 9. Gruenwald P. Normal and abnormal detachment of body and gut from the blastoderm in the chick embryo, with remarks on the early development of the allantois. J Morph 1941; 69:83-125. 10. Hamburger V, Hamilton HL. A series of normal stages in the development of the chick embryo. J Morph 1951; 88:49-92. 11. Romanoff AL, The Avian Embryo. 1960, New York: The Macmillan Company. 1305. 12. Le Douarin N. Etude experimentale de l’organeogenese du tube difestif et du foie chez l’embryon de poulet. Bull Biol France, Belg 1964; 98:533-676. 13. Haffen K, Lacroix B, Kedinger M, Simon-Assman PM. Inductive properties of fibroblastic cell cultures derived from rat intestinal mucosa on epithelial differentiation. Differentiation 1983; 23:226-233. 14. Haffen K, Keddinger M, Simon-Assman P. Mesenchyme-dependent differentiation of epithelial progenitor cells in the gut. J Pediatric Gastroenterology and Nutrition 1987; 6:14-23. 15. Yasugi S. Role of epithelial-mesenchymal interactions in differentiation of epithelium of vertebrate digestive organs. Develop Growth & Differ 1993; 35:1-9. 16. Kedinger M, Simon-Assman PM, Bouziges F, Arnold C, Alexandre E, Haffen K. Smooth muscle actin expression during rat gut development and induction in fetal skin fibroblastic cells associated with intestinal embryonic epithelium. Differentiation 1990; 43:87-97. 17. Sumiya M. Differentiation of the digestive tract epithelium of the chick embryo cultured in vitro enveloped in a fragment of vitelline membrane in the absence of mecenchyme. Roux’s Archiv 1976; 197:1-17. 18. Aufderheide E, Ekblom P. Tenascin during gut development: Appearance in the mesenchyme, shift in molecular forms, and dependence on epithelial-mesenchymal interactions. J Cell Biol 1988; 107:2341-2349. 19. Kedinger M, Simon-Assman P, Bouziges F et al. Epithelial-mesenchymal interactions in intestinal epithelial differentiation. Scand J Gastroenterol 1988; 23:62-69.

Epithelial-Mesenchymal Interactions in Gastrointestinal Development

197

20. Ishizuya-Oka A. Electron microscopical study of self-differentiation potency in the chick embryonic endoderm cultured in vitro. Roux’s Arch. Dev Biol 1983; 192:171-178. 21. Ishizuya-oka A, Mizuno T. Intestinal cytodifferentiation in vitro of chick stomach endoderm induced by the duodenal mesenchyme. J Embryol exp Morph 1984; 82:163-176. 22. Hayashi K, Agata K, Mochii M et al. Molecular cloning and the nucleotide sequence of cDNA for embryonic chicken pepsinogen: Phylogenetic relationship with prochymosin. J Biochem 1988; 103:290-296. 23. Hayashi K, Yasugi S, Mizuno T. Pepsinogen gene transcription induced in heterologous epithelial-mesenchymal recombinations of chicken endoderms and glandular stomach mesenchyme. Development 1988; 103:725-731. 24. Takiguchi K, Yasugi S, Mizuno T. Developmental changes in the ability to express embryonic pepsinogen in the stomach epithelia of chick embryos. Roux’s Arch. Dev Biol 1988; 197:56-62. 25. Fukamachi H, Takayama S. Epithelial-mesenchymal interaction in differentiation of duodenal epithelium of fetal rats in organ culture. Experientia 1980; 36:335-336. 26. Fukamachi H, Mizuno T, Takayama S. Epithelial-Mesenchymal interactions in differentiation of stomach epithelium in fetal mice. Anat Embryol 1979; 157:151-160. 27. Takiguchi K, Yasugi S, Mizuno T. Gizzard epithelium of chick embryos can express embryonic pepsinogen antigen, a marker protein of proventriculus. Roux’s Arch Dev Biol 1986; 195:475-483. 28. Yasugi S. Differentiation of allantoic endoderm implanted into the presumptive digestive area in avian embryos. A study with organ-specific antigens. J Embryol exp Morph 1984; 80:137-153. 29. Yasugi S, Matsushita S, Mizuno T. Gland formation induced in the allantoic and smallintestinal endoderm by the proventricular mesenchyme is not coupled with pepsinogen expression. Differentiation 1985; 30:47-52. 30. Yasugi S, Takeda H, Fukuda K. Early determination of developmental fate in preseumtive intestinal endoderm of the chicken embryo. Devel Growth & Diff 1991; 33:235-241. 31. Fukuda K, Ishii Y, Saiga H et al. Mesenchymal regulation of epithelial gene expression in developing avian stomach: 5' flanking region of pepsinogen gene can mediate mesenchymal influence on its expression. Development 1994; 120:3487-3495. 32. Duluc I, Freund J-N, Leberquier C et al. Fetal endoderm primarily holds the temporal and positional information required for mammalian intestinal development. J Cell Biol 1994; 126:211-221. 33. Mathan M, Hermos JA, Trier JS. Structural features of the epithelio-mesenchymal interface of rat duodenal mucosa during development. J Cell Biol 1972; 52:577-588. 34. Fukushige T, Schroeder DF, Allen FL et al. Modulation of gene expression in the embryonic digestive tract of C. elegans. Dev Biol 1996; 178:278-288. 35. Tepass U, Hartenstein V. Epithelium formation in the Drosophila midgut depends on the interaction of endoderm and mesoderm. Development 1994; 120:579-590. 36. Takiguchi-Hayashi K, Yasugi S. Transfilter analysis of the inductive influence of proventricular mesenchyme on stomach epithelial differentiation of chick embryos. Roux’s Arch Dev Biol 1990; 198:460-466. 37. Simon-Assman P, Bouziges F, Arnold C et al. Epithelial-mesenchymal interactions in the production of basement membrane components in the gut. Development 1988; 102:339-347. 38. Schmidt JW, Piepenhagen PA, Nelson WJ. Modulation of epithelial morphogenesis and cell fate by cell-to-cell signals and regulated cell adhesion. Sem Cell Biol 1993; 4:161-173. 39. Kedinger M, Bouziges F, Simon-Assmann P et al. Influence of cell interactions on intestinal brush border enzyme expression, in Highlights Modern Biochemistry, A Kotyz, J Skoda, V Paces et al, Editors. 1989, VSP International, Zeist. p. 1103-1112. 40. Hahn U, Stallmach A, Hahn EG et al. Basement membrane components are potent promoters of rat intestinal epithelial cell differentiation in vitro. Gastroenterology 1990; 989:322-335. 41. Simon-Assmann P, Kedinger M. Heterotypic cellular cooperation in gut morphogenesis and idfferentiation. Sem Cell Biol 1993; 4:221-230.

198

Molecular Basis of Epithelial Appendage Morphogenesis

42. Weiser MM, Sykes DE, Killen PD. Rat intestinal basement membrane synthesis, epithelial versus nonepithelial contributions. Lab Invest 1990; 62:325-330. 43. Simo P, Bouziges F, Lissitzky JC et al. Dual and asynchronous desposition of laminin chains at the epithelial-mesenchymal interface in the gut. Gastroenterology 1992; 102:1835-1845. 44. Hayashi K. Thinning of the basement membrane and localized cell proliferation during gland formation of the chick proventriculus. Develop Growth & Differ 1987; 29:285-295. 45. Beaulier JF, VCachon PH, Chartrand S. Immunolocalization of extracellular matrix components during organogenesis in the human small intestine. Anat Embryol 1991; 183:363-369. 46. Takiguchi-Hayashi K, Yasugi S. Localized distribution of a novel mesenchyme-specific antigen in developing chick digestive organs: Comparison with the distribution of fibronectin, laminin and tenascin. Roux’s Arch Dev Biol 1991; 200:113-116. 47. Simpson P. Maternal-zygotic gene interactions during formation off the dorsoventral pattern in Drosophila embryo. Genetics 1983; 105:615-632. 48. Grau Y, Carteret C, Simpson P. Mutations and chromosomal rearrangements affecting the expression of snail, a gene involved in embryonic patterning in Drosophila melanogaster. Genetics 1984; 108:347-360. 49. Tepass U, Gruszynski-DeFeo E, Haap TA et al. Shotgun encodes Drosophila E-cadherin and is preferentially required during cell rearrangement in the neurectoderm and other morphgenetically active epithelia. Genes Dev 1996; 10:672-685. 50. Duncan IM. The bithorax complex. Ann Rev Genet 1987; 21:285-319. 51. Kaufman TC, Seeger MA, Olsen G. Molecular and genetics organization of the Antennapedia gene complex of Drosophila melanogaster. Adv Genet 1990; 27:309-362. 52. Lawrence PA, Morata G. Homeobox genes: Their function in Drosophila segmentation and pattern formation. Cell 1994; 78:181-189. 53. Tremml G, Bienz M. Homeotic gene expression in the visceral mesoderm of Drosophila embryos. EMBO J 1989; 8:2677-2685. 54. Immergluck K, Lawrence PA, Bienz M. Induction across germ layers in Drosophila mediated by a genetic cascade. Cell 1990; 62:261-268. 55. Panganiban GF, Reuter R, Scott MP et al. A Drosophila growth factor homolog, decapentaplegic, regulates homeotic gene expression within and across germ layers during midgut morphogenesis. Development 1990; 100:1041-1050. 56. Reuter R, Panganigan GEF, Hoffmann FM et al. Homeotic genes regulate the spatial expression of putative growth factors in the visceral mesoderm of Drosophila embryos. Development 1990; 110:1031-1040. 57. Bienz M. Homeotic genes and the positional signalling in the Drosophila viscera. Trends Genet 1994; 10:22-26. 58. Hopper S, Bienz M. Specification of a single cell bype by a Drosophila homeotic gene. Cell 1994; 76:689-702. 59. Mathies LD, Kerridge S, Scott MP. Role of the teashirt gene in Drosophila midgut morphogenesis: secreted proteins mediate the actrion of homeotic genes. Development 1994; 120:2799-2809. 60. Mann RS, Abu-Shaar M. Nuclear import of the homeodomain protein Extradenticle in response to Wg and Dpp signalling. Nature 1996; 383:630-633. 61. Bate M, Arias AM, eds. The Development of Drosophila melanogaster. 1 ed. 1993, Cold Spring Harbor Laboratory Press. 1558. 62. MacDonald PM, Struhl G. A molecular gradient in early Drosophila embryos and its role in specifying the body pattern. Nature 1986; 324:537-545. 63. Pankratz MJ, Hoch M. Control of epithelial morphogenesis by cell signalling and integrin molecules in Drosophila foregut. Development 1995; 121:1885-1898. 64. Kaur S, Singh G, Stock JL et al. Dominant mutation of the murine Hox-2.2 gene results in developmental abnormalities. J Exp Zoo 1992; 264:323-36. 65. Dolle P, Izpisua-Belmonte JC, Brown J et al. Hox genes and the morphogenesis of the vertebrate limb. [Review]. Prog Clin Biol Res 1993; 11-20. 66. McGinnis W, Krumlauf R. Homeobox genes and axial patterning. Cell 1992; 68:283-302.

Epithelial-Mesenchymal Interactions in Gastrointestinal Development

199

67. Krumlauf R. Hox genes in vertebrate development. Cell 1994; 78:191-201. 68. Benson GV, Lim H, Paria BC et al. Mechanisms of reduced fertility in HoxA10 mutant mice: Uterine homeosis and loss of maternal HoxA10 expression. Development 1996; 122:2687-2696. 69. Hsieh-Li HM, Witte DP, Weinstein M et al. HoxA11 structure, extensive antisense transcription, and function in male and female fertility. Development 1995; 121:1373-1385. 70. Manley NR, Capecchi MR. The role of HoxA3 in mouse thumus and thyroid development. Development 1995; 121:1989-2003. 71. Dolle P, Izpisua-Belmonte JC, Brown JM et al. HOX-4 genes and the morphogenesis of mammalian genitalia. Genes Dev 1991; 5:1767-7. 72. Izpisua-Belmonte JC, Falkenstein H, Dolle P et al. Murine genes related to the Drosophila AbdB homeotic genes are sequentially expressed during development of the posterior part of the body. EMBO J 1991; 10:2279-89. 73. Dolle P, Dierich A, LeMeur M et al. Disruption of the Hoxd13 gene induces localized heterchrony leading to mice with neotenic limbs. Cell 1993; 75:431-441. 74. Roberts DJ, Johnson RL, Burke AC et al. Sonic Hedgehog is an endodermal signal inducing BMP-4 and Hox genes during induction and regionalization of the chick hindgut. Development 1995; 121:3163-3174. 75. Yokouchi Y, Sakiyama J, Kuroiwa A. Coordinated expression of Abd-B subfamily genes of the HoxA cluster in the developing digestive tract of the chick embryo. Dev Biol 1995; 169:76-89. 76. Tabin CJ. Why we have (only) five fingers per hand: hox genes and the evolution of paired limbs. [Review]. Development 1992; 116:289-96. 77. Nelson CE, Morgan BA, Burke AC et al. Analysis of Hox gene expression in the chick limb bud. Development 1996; 122:1449-1466. 78. Wolgemuth DJ, Behringer RR, Mostoller MP et al. Transgenic mice overexpressing the mouse homoeobox-containing gene Hox-1.4 exhibit abnormal gut development. Nature 1989; 337:464-7. 79. Behringer RR, Crotty DA, Tennyson VM et al. Sequences 5' of the homeobox of the Hox1.4 gene direct tissue-specific expression of lacZ during mouse development. Development 1993; 117:823-33. 80. Gaunt SJ, Krumlauf R, Duboule D. Mouse homeo-genes within a subfamily, Hox-1.4, -2.6 and -5.1, display similar anteroposterior domains of expression in the embryo, but show stage- and tissue-dependent differences in their regulation. Development 1989; 107:131-41. 81. Galliot B, Dolle P, Vigneron M et al. The mouse Hox-1.4 gene: primary structure, evidence for promoter activity and expression during development. Development 1989; 107:343-59. 82. Gaeda AM, Gaunt SJ, Azzawi M et al. Sequence and embryonic expression of the murine Hox-3.5 gene. Development 1992; 116:497-506. 83. Boulet AM, Capecchi MR. Targeted disruption of hoxc-4 causes esophageal defects and vertebral transformations. Dev Biol 1996; 177:232-249. 84. Pollock RA, Jay G, Bieberich CJ. Altering the boundaries of Hox3.1 expression: Evidence for antipodal gene regulation. Cell 1992; 71:911-23. 85. Kondo T, Dolle P, Zakany J et al. Function of Posterior HoxD genes in the morphogenesis of the anal sphincter. Development 1996; 122:2651-2659. 86. Fromental-Ramain C, Warot X, Lakkaraju S et al. Specific and redundant functions of the paralogous HoxA9 and HoxD9 genes in forelimb and axial skeleton patterning. Development 1996; 122:461-472. 87. Horan GSB, Ramirez-Solis R, Featherstone MS et al. Compound mutants for the paralogous hoxa-4, hoxb-4, and hoxd-4 genes show more complete homeotic transformations and a dose -dependent increase in the number of vertebrae transformed. Genes Dev 1995; 9:1667-1677. 88. Grobstein C. Induction interction in the development of the mouse metanephros. J Exp Zool 1955; 130:319-340.

200

Molecular Basis of Epithelial Appendage Morphogenesis

89. Grobstein C. Trans-filter induction of tubules in mouse metanephrogenic mesenchyme. Exp Cell Res 1956; 10:424-440. 90. Small KM, Potter SS. Homeotic transformations and limb defects in Hox A11 mutant mice. Genes Dev 1993; 7:2318-28. 91. Davis AP, Witte DP, Shieh-li HM et al. Absence of radius and ulna in mice lacking hoxa11 and hoxd-11. Nature 1995; 375:791-795. 92. Averof M, Akam M. Hox genes and the diversification of insect and crustacean body plans. Nature 1995; 376:420-423. 93. van der Hoeven F, Zakany J, Duboule D. Gene transpositions in the HoxD complex reval a hierarchy of regulatory controls. Cell 1996; 85:1025-1035. 94. Schummer M, Scheurlen I, Schaller C et al. HOM/HOX homeobox genes are present in hydra (Chlorohydra viridissima) and are differentially expressed during regeneration. EMBO J 1992; 11:1815-1823. 95. Balavoine G, Talford MJ. Indentification of planarian homeobox sequences indicates the antiquity of most Hox/homeotic gene subclasses. Proc Natl Acad Sci USA 1995; 92:7227-7231. 96. Riddle RD, Johnson RL, Laufer E et al. Sonic hedgehog mediates the polarizing activity of the ZPA. Cell 1993; 75:1401-16. 97. Johnson RL, Laufer E, Riddle RD et al. Ectopic expression of Sonic hedgehog alters dorsalventral patterning of somites. Cell 1994; 79:1165-1173. 98. Fan CM, Tessier-Lavigne M. Patterning of mammalian somites by surface ectoderm and notochord: Evidence for sclerotome induction by a hedgehog homolog. Cell 1994; 79:1175-1186. 99. Echelard Y, Epstein DJ, St-Jacques B et al. Sonic hedgehog, a member of a family of putative secreted signaling molecules, is implicated in the regulation of CNS polarity. Cell 1993; 75:1417-1430. 100. Krauss S, Concordet J-P, Ingham PW. A functionally conserved homolog of the Drosophila segment polarity gene hh is expressed in tissues with polarizing activity in zebrafish. Cell 1993; 75:1431-1444. 101. Roelink H, Augsburger A, Heemskerk J et al. Floor plate and motor neuron induction by vhh-1, a vertebrate homolog of hedgehog expressed by the notochord. Cell 1994; 76:761-775. 102. Laufer E, Nelson CE, Johnson RL et al. Sonic hedgehog and Fgf-4 act through a signaling cascade and feedback loop to integrate growth and patterning of the developing limb bud. Cell 1994; 79:993-1003. 103. Niswander L, Jeffrey S, Martin GR et al. A positive feedback loop coordinates growth and patterning in the vertebrate limb. Nature 1994; 371:609-612. 104. Marigo V, Roberts DJ, Lee SMK et al. Cloning, expression, and chromosomal location of SHH and IHH, two human homologues of the Drosophila segment polarity gene hedgehog. Genomics 1995; 28:44-51. 105. Chiang C, Litingtung Y, Lee E et al. Cyclopia and defective axial patterning in mice lacking Sonic hedgehog gene function. Nature 1996; 383:407-413. 106. Bitgood MJ, McMahon AP. Hedgehog and Bmp genes are coexpressed at many diverse sites of cell-cell interaction in the mouse embryo. Dev Biol 1995; 172:126-138. 107. Bellusci S, Henderson R, Winnier G et al. Evidence from normal expression and targeted misexpression that Bone Morphogenetic Protein-4 (Bmp-4) plays a role in mouse embryonic lung morphogenesis. Development 1996; 122:1693-1702. 108. Winnier G, Blessing M, Labhosky PA et al. Bone morphogenetic protein-4 is required for mesoderm formation and patterning in the mouse. Genes Dev 1995; 9:2103-2116. 109. Kaestner KH, Lee K-H, Schlondorff J et al. Six members of the mouse forkhead gene family are developmentally regulated. Proc Natl Acad Sci USA 1993; 90:7628-7631. 110. Monaghan AP, Kaestner KH, Grau E et al. Postimplantation expression patterns indicate a role for the mouse forkhead/HNF-3 a, b, g genes in determination of the definitive endoderm, chordamesoderm and neuroectoderm. Development 1993; 119:567-578.

Epithelial-Mesenchymal Interactions in Gastrointestinal Development

201

111. Sasaki H, Hogan BLM. Differntial expression of multiple fork head related genes during gastrulation and axial pattern formation in the mouse embryo. Development 1993; 118:47-59. 112. Ang S-L, Wierda A, Wong D et al. The formation and maintenance of the definitive endoderm lineage in the mouse: Involvement of HNF3/forkhead proteins. Development 1993; 119:1301-1305. 113. Sasaki H, Hogan BLM. HNF-3 as a regulator of floor plate development. Cell 1994; 76:103-115. 114. Levin M, Johnson RL, Stern CD et al. A molecular pathway determining left-right asymmetry in chick embryogenesis. Cell 1995; 82:803-814. 115. Buchberger A, Pabst O, Brand T et al. Chick NKx2.3 represents a novel family member of vertebrate homologues to the Drosophila homeobox gene tinman: Differential expression of cNKx2.3 and cNKx2.5 during heart and gut development. Mech Devel 1996; 56:151-163. 116. Tissier-Seta J-P, Mucchielli M-L, Mark M et al. Barx1, a new mouse homeodomain transcription factor expressed in cranio-facial ectomesenchyme and the stomach. Mech Dev 1995; 51:3-15. 117. Chuong C-M, Jiang T-J, Yin E et al. cDCC (chicken homologue to a gene deleted in colorectal cardinoma) is an epithelial adhesion molecule expressed in the basal cells and involved in epithelial-mesenchymal interaction. Dev Biol 1994; 164:383-397. 118. Hirai Y, Takebe K, Takashina M et al. Epimorphin: A mesenchymal protein essential for epithelial morphogenesis. Cell 1992; 69:471-481. 119. Andermarcher E, Surani MA, Gherardi E. Co-expression of the HGF/SF and c-met genes during early mouse embryogenesis precedes reciprocal expression in adjacent tissues during organogenesis. Dev Gen 1996; 18:254-266. 120. Muller TS, Ebensperger C, Neubuser A et al. Expression of avian Pax1 and Pax9 is intrinsically regulated in the pharyngeal endoderm, but depends on environmental influences in the paraxial mesoderm. Dev Biol 1996; 178:403-417. 121. Frumkin A, Pillemer G, Haffner R et al. A role for CdxA in gut closure and intestinal epithelial differentiation. Development 1994; 120:253-263. 122. Doll U, Niessing J. Continued expression of the chicken caudal homologue in endodermally derived organs. Dev Biol 1993; 156:155-163. 123. James R, Kazenwadel J. Homeobox gene expression in the intestinal epithelium of adult mice. J Biol Chem 1991; 266:3246-3251. 124. Meyer BI, Gruss P. Mouse Cdx-1 expression during gastrulation. Development 1993; 117:191-203. 125. Frumkin A, Rangini A, Ben-Yehuda A et al. A chicken caudal homologue, CHox-cad, is expressed in the epiblast with posterior localization and in the early endodermal lineage. Development 1991; 112:207-219. 126. Freund J-N, Boukamel R, Benazzouz A. Gradient expression of Cdx along the rat intestine throughout postnatal development. FEBS 1992; 314:163-166. 127. Frumkin A, Haffner R, Shapira E et al. The chicken CdxA homeobox gene and axial positioning during gastrulation. Development 1993; 118:553-562. 128. Suh E, Chen L, Tayler J et al. A homeodomain protein related to caudal regulated intestinspecific gene transcription. Mol Cell Biol 1994; 14:7340-7351. 129. James R, Erler T, Kazenwadel J. Structure of the murine homeobox gene cdx-2. J Biol Chem 1994; 269:15229-15237. 130. Gamer LW, Wright CVE. Murine Cdx-4 bears striking similarities to the Drosophila caudal gene in its homeodomain sequence and early expression pattern. Mech Devel 1993; 43:71-81. 131. Jonsson J, Carlsson L, Edlund T et al. Insulin-promoter-factor 1 is required for pancreas development in mice. Nature 1994; 371:606-609. 132. Ahlgren U, Johsson J, Edlund H. The morphogenesis of the pancreatic mesenchyme is uncoupled from that of the pancreatic epithelium. Development 1996;122:1409-1416. 133. Offield MF, Jetton TL, Labosky PA et al. PDX-1 is required for pancreatic outgrowth and differentiation of the rostral duodenom. Development 1996; 122:983-995.

CHAPTER 11

Endodermal Appendage Formation: Morphogenetic Mapping of Dorso-Ventral Patterning of the Anterior Foregut and Development of Lung and Thyroid Primordia Parviz Minoo, Shioko Kimura and Robert deLemos

Introduction

T

he endodermal cells of the gut serve as progenitors for the origin of many diverse tissues including pancreas, liver, thyroid and the lung. The primordium of each tissue is thought to emerge as an endodermal appendage consisting of a small group of cells with a specific developmental fate. One mechanism by which the progenitor of each tissue can be distinguished from all others is by creating molecularly discernible regional differences along the gut axis. Regionalization must include determination of the exact spatial and temporal coordinates of each primordial appendage and its relationship with other neighboring structures. Once specified, morphogenesis of each progenitor into a functionally mature organ requires further regulation, which includes cellular proliferation, migration, apoptosis and differentiation. How is such a precise, multidimensional plan implemented? In the most anterior portion of the foregut, within a relatively narrow zone which ultimately forms the pharynx, is the site where at least three structures—the esophagus, trachea and the thyroid glands—come into close proximity. Normal morphogenesis of this zone is accomplished by pattern formation, through which the precise spatial relationship of the three tissues along the dorso-ventral and the antero-posterior axes of the foregut is established. The goal of this chapter is to present recent experimental data regarding the role of a specific transcriptional factor, Nkx-2.1, in pattern formation in the anterior foregut and morphogenesis of thyroid and lung tissues. Transcriptional regulation by Nkx-2.1 can be further used as a paradigm for regulation of morphogenesis in other endodermally derived tissue appendages.

Progenitor Field Specification Lessons from Early Specification of Body Plans in Invertebrates The critical question of how a group of cells is specified to form the progenitor of a specialized organ, and the mechanisms which govern its growth and morphogenesis, remains largely unanswered. However, to develop an operational concept of how this process Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

204

Molecular Basis of Epithelial Appendage Morphogenesis

may take place we can take advantage of the fact that fundamental themes in development are evolutionarily conserved, and the concepts derived from genetic and molecular studies in morphogenesis of Caenorhabditis, Drosophila and recently zebrafish (Danio rerio) may have analogous counterparts in signaling and specification of tissue progenitors during organogenesis in mammals. It is now clear that the body plan of metazoans is established by creating molecular differences within a seemingly homogeneous mass of cells that make up the early embryo. In the fruit fly Drosophila melanogaster embryos, establishment of the head-tail axis is accomplished by products of two genes, nanos and bicoid, whose mRNAs are expressed from the maternal genome and packaged into the oocyte at the anterior and posterior poles of the egg respectively.1-3 Upon fertilization, translation of the maternal mRNAs produces gradients of the two proteins in the early embryo. The concentration gradient (highest in the anterior compartment) of bicoid, a transcriptional factor, activates the zygotic expression of another transcriptional factor, hunchback.4,5 Gradient distribution of hunchback (again highest at the anterior pole) in turn activates a number of other transcriptional factors. Different concentrations of hunchback activate different sets of genes along the anteroposterior axis of the embryo, leading to the establishment of a segmented and “polarized” structure. Once the segmented pattern of the body plan is laid down, the identity of the segments is determined by which combination of other transcriptional regulators encoded by “homeotic genes” are selectively activated and expressed in each segment.4 A major distinction, however, between the fly embryo and that of most animals is that the fly embryo is a syncytium composed of nuclei sharing the same cytoplasm, whereas other embryos consist of well defined cells. Therefore, whereas in the fly a gradient of transcriptional factors can directly activate gene batteries in individual or groups of nuclei, early embryogenesis in vertebrates requires signaling mechanisms, in the form of soluble factors and cell surface receptors, through which the localized developmental cues are transmitted from one cell to its neighbors. The latter process is the phenomenon of cell-cell interactions which is covered elegantly in chapter 10 using gut development as a paradigm. It is generally accepted that during later periods of embryogenesis the progenitors of various organs are specified in the form of what classically has been termed an anlage. Formation of an anlage, or as Davidson called it the “progenitor field”,6 can occur through a signaling mechanism that specifies the position of this structure, the cells of which the structure will consist, and its boundaries with respect to the overall body plan. Because the cells within a progenitor field can be distinguished from other fields by the array of specific genes they express, the target of the signal that initiates a progenitor field is likely a transcriptional regulator. In this scheme, the signaling must always lie upstream of the genes encoding the transcriptional regulators.6 Therefore, the sequence of events during organogenesis will likely consist of: Signaling → Transcriptional regulator → Morphogenesis The signals which specify a progenitor “field” in vertebrate organogenesis are unknown. However, once specified, the cells demarcated to form a given tissue are guided by a morphogenetic plan otherwise known as pattern formation, through which specific structures of the correct shape, size, position and orientation are generated during embryogenesis. In its broadest definition, pattern formation describes and is not only critical for the establishment of the initial body plan of the early embryo but is also applicable as a conceptual framework for the way in which every part of the embryo develops through morphogenesis. Wolpert7,8 introduced the useful concept of “positional information” to describe how pattern formation could be accomplished by the individual cell. According to this scheme, a cell parameter exists, called positional value, which is related to a cell’s position in the coor-

Endodermal Appendage Formation

205

dinates of the developing system. Within this coordinate system the cells have a given positional value and the behavioral decisions they make, be it proliferation, migration or differentiation, are based on their interpretation of this value. Assignment of positional information could be accomplished through signal gradients which can be passed from one cell to the next.

Hox Genes Are the Target of Signals That Specify and Regulate Pattern Formation Homeotic genes, HOMs, were first described in Drosophila based on genetic evidence of their mutated alleles to cause “homeosis” or replacement of one body part for another.9,10 There is now molecular and genetic evidence that the two clusters of homeotic genes within the Antennapedia (ANT-C) and bithorax (BX-C) complexes, collectively known as the HomC locus, function in specification of segmental identity.11,12 Loss of function mutations in these loci result in transformation of one segment into an adjacent identity. All genes within the Hom-C complex encode proteins which have an evolutionarily highly conserved, sequence-specific, DNA-binding domain known as the “homeodomain”.13 There is evidence that several of these proteins function as transcriptional factors.14 Therefore, the function of the Hom genes may be in activation of target genes necessary within a given segment for differentiation of its cells and establishment of phenotypic identity. Largely based on low stringency hybridization, homologues of Drosophila Hom genes, called Hox, have been isolated and characterized in vertebrates. Four clusters of Hox genes have been identified. The clusters are highly conserved both in structure and genetic organization.15,16 The physical order of the genes within a cluster is colinear with their temporal appearance and the antero-posterior boundary of expression along the embryonic axis.15 Other Hox genes, whose homeodomain structures are divergent from those found in the four clusters, are also expressed in tissues. Many of the Hox genes essential for morphogenetic processes that lead to organogenesis fall in this category. The current concept of how the products of homeobox-containing genes act during development is based on what is known genetically and biochemically about their counterparts in Drosophila. The Hox genes have been found to be expressed in segmental units in the hindbrain or rhombomeres and paraxial mesoderm.17 However, expression of Hox genes in nonsegmented tissues suggests that their gene products may be important in the specification of regional identity along embryonic axes. In summary, members of transcriptional regulators, and particularly those belonging to the homeodomain class, are likely to control regional specification of progenitor fields. Hox genes are downstream of the signaling systems which initiate the process of regional specification. However, since their expression can control inductive interactions that divide and distinguish morphogenetic fields, they could also act upstream of the signal systems.

Pattern Formation in the Anterior Foregut Can these concepts be applied to pattern formation in the anterior foregut and to morphogenesis of endodermal appendages which originate from this tissue? The primordium of the lung and the thyroid are endodermal appendages that are derived at around E9 to E10 from specific cells within a narrow zone of the anterior foregut.18-20 Formation of each primordium occurs with correct spatial and temporal specificity with respect to the overall body plan. What signal(s) dictate these parameters is unknown. From a simplistic point of view, a group of cells along the gut axis is set aside by virtue of a given and distinct set of genes activated in response to an initial signal. The activated genes are necessary for regulating pattern formation in the thyroid and the lung through which the processes of proliferation/migration and differentiation are integrated to form a structure with the correct shape,

206

Molecular Basis of Epithelial Appendage Morphogenesis

size and orientation. In recent years it has become clear that organogenesis, in much the same way as early embryogenesis, is accomplished by sequential activation of spatially and temporally regulated genes. In each tissue, a hierarchy of regulatory genes constitutes a developmental map, based on which determination and subsequent morphogenesis of that tissue are implemented.

Regionalization of the Gut by Spatially-restricted Expression of the Hnf-3/forkhead Gene Family The underlying mechanisms that govern normal patterning of the anterior foregut have not been elucidated. In constructing a developmental map, one approach is to correlate the time of activation and spatial distribution of specific regulatory genes with the onset of morphogenetic events. Following the theme of Signal → Transcriptional factors → Morphogenesis, many investigators have sought to correlate the expression of transcriptional factors with morphogenetic events. As a result, the hepatocyte nuclear factor, HNF-3/ forkhead gene family has been identified amongst the first regulatory genes whose expression occurs in the definitive endoderm during early embryogenesis.21,22 HNF-3/forkhead genes exhibit striking structural and functional similarities with members of the Drosophila forkhead (fkh) gene family.23-26 Importantly, expression of HNF-3/forkhead genes persists in endodermally derived adult structures.27 The lung and the thyroid primordia are specified within the boundaries of expression of HNF-3α and HNF-3β.27 HNF-3α and HNF-3β mRNA are detected as early as the time when the esophagus and laryngo-tracheal groove begin to differentiate (mouse E9.5). Both genes are expressed in fully differentiated adult bronchial epithelium27 and regulate the lung specific promoters CC10 and SP-B.28,29 Similarly, in the thyroid, HNF3-β participates in the transcriptional regulation of the promoter of the thyroperoxidase gene.30 Transcripts from another member, HNF-3γ, are detectable during hindgut differentiation and persist through liver and stomach morphogenesis. Thus regionally-specific expression of HNF-3/forkhead genes may constitute a molecular axial code which specifies the progenitors of the structures that emerge from the gut endoderm. Whether activation of the HNF-3 gene family is necessary or sufficient for specification of the lung and thyroid primordia remains to be experimentally determined. In this regard, disruption of one member of this gene family, HNF-3β, by homologous recombination results in embryonic lethality where the normal notochord and the node fail to develop.31,32 In these mutants, foregut morphogenesis is arrested at an early stage during embryogenesis, which in turn results in the absence of lung, thyroid and other foregut derivatives. Because of its early lethal phenotype, this mutant has not been informative in deciphering further steps in morphogenesis of the anterior foregut and emergence of specific foregut-derived tissue primordia.

Further Regionalization Nkx-2.1 Expression Distinguishes Thyroid and Lung Primordia from Other Endodermally Derived Tissues Subsequent to establishment of the foregut, a homeodomain transcriptional factor, Nkx-2.1 (otherwise known as TTF-1 or T/ebp), is expressed in two of the many appendages which emerge from the gut endoderm, the thyroid and the lung.33 Based on its ability to bind and direct transcription from thyroid specific gene promoters, Nkx-2.1 was first purified by Civitareale et al.34 Subsequently, cDNA clones from rat and mouse thyroid cDNA libraries were isolated which contained the full length open reading frame.35,36 Nkx-2.1 is encoded by a single gene in the rat, mouse and the human.37-39 The encoded protein contains a divergent and novel DNA binding domain with considerable sequence similarity to

Endodermal Appendage Formation

207

the Drosophila NK-2 class of homeodomains.40 The homeodomain is sufficient for the DNA binding activity of the protein. The binding specificity of Nkx-2.1 homeodomain is distinct from that of the Antennapedia class.41 The crucial difference appears to be in the motif 5'-TAAT-3', which is recognized by Antennapedia, and 5'-CAAG-3' preferentially bound by Nkx-2.1. The CAAG motif is present in the promoter regions of both thyroid and lung specific genes.29,42,30 Expression of Nkx-2.1 occurs at the onset of lung and thyroid morphogenesis but is also detectable in select parts of the brain.33 In the lung, Nkx-2.1 is expressed in all epithelial cells early in pulmonary morphogenesis, but becomes progressively confined to specific epithelial cell types such as alveolar type II and Clara cells. In the proximal respiratory compartments, Nkx 2.1 expression is not detectable in tracheal and primary bronchial epithelial cells. In vitro data suggest that both Nkx-2.1 and HNF-3 have the ability to bind to and activate transcription of pulmonary specific genes.29,42,43 In E9.5 day mouse embryos, Nkx-2.1 is also expressed in the primordium of the thyroid gland which, like that of the lung, originates from the ventral wall of the anterior foregut.33 The thyroid primordium is formed from a group of endodermal cells which appear just above the dilated aortic sac at around mouse embryonic day 9 to 10.19 In E11 mouse embryos, when the thyroid diverticulum assumes the shape of a vesicle, Nkx-2.1 expression can be detected on the ventral edge and not on the posterior remnant of the thyroglossal duct. By day 17 of gestation, Nkx-2.1 expression becomes restricted to the cords of the epithelial cells which are still arranged in an irregular interdigitating pattern.33

Nkx-2.1 Is Required for Normal Dorso-Ventral Patterning of the Anterior Foregut Homologous recombination was recently used to generate a strain of mice that carries a targeted disruption of the Nkx-2.1 locus.44 Nkx-2.1 (-/-) embryos die shortly after birth due to respiratory insufficiency. Functional deletion of Nkx-2.1 is accompanied by phenotypic abnormalities in three structures, all of which originate from the anterior foregut. These include dysfunctional hypoplastic lungs, absence of thyroid tissue and a failure in septation between the tracheal and esophageal lumens. The latter phenotype is strikingly similar to a clinical condition in human newborns known as tracheo-esophageal (TE) fistula.45 The underlying mechanism responsible for this debilitating and sometimes lethal condition is entirely unknown. TE fistulas result from a failure in septation between tracheal and the esophageal lumens, both of which are derived from the foregut endoderm. In Nkx-2.1 (-/-) mouse embryos, the pharynx is connected through a single lumen to the stomach. The two primary (main-stem) bronchi emerge bilaterally from this shared luminal structure, connecting it to two markedly hypoplastic lungs. The more cranial portion of the common lumen shows phenotypic characteristics of both trachea and esophagus, including rings of cartilage. However, the composition and the number of tracheal cartridges in Nkx-2.1 (-/-) mutants deviate significantly from those in wild type mice. During normal development, tracheal cartilages are derived from splanchnic mesenchyme through interactions with the developing lung epithelium.46 The abnormalities in the number and morphology of the tracheal cartilage in Nkx-2.1 (-/-) mutants suggest that the absence of Nkx-2.1 disrupts normal epithelial-mesenchymal interactions. The overall observations regarding the phenotypic characteristics of the Nkx-2.1 (-/-) mouse embryos are consistent with failure of the anterior foregut to undergo normal compartmentalization into distinct tracheal and esophageal structures. Therefore, it would appear that absence of a functional Nkx-2.1 allele leads to abrogation of normal morphogenetic pathways which divide the foregut into ventral and dorsal compartments.

208

Molecular Basis of Epithelial Appendage Morphogenesis

The relationship between the expression of Nkx-2.1 and dorso-ventral compartmentalization of the anterior foregut has been addressed.47 Consistent with the observed phenotype in the knockout mouse, Nkx-2.1 expression is detectable only in a select group of endodermal cells surrounding the ventral wall of the anterior foregut in wild type mouse embryos at E9.5, the time of emergence of the laryngo-tracheal diverticulum (respiratory primordium). Nkx-2.1 expression is not detectable in the endodermal cells surrounding the dorsal wall of the anterior foregut, the site of emergence of the esophagus. Therefore, the expression pattern of Nkx-2.1 defines the dorso-ventral boundary within the anterior foregut thus clearly distinguishing the tracheal and pulmonary primordium from that of the esophagus. The mechanism by which Nkx-2.1 is selectively activated in the ventral half of the anterior foregut remains to be deciphered. In Drosophila, examination of NK-2 gene expression in mutant embryos with dorso-ventral defects has identified a number of genes which affect the expression of NK-2.48 These studies show that NK-2 is activated by dorsal49-51 in the ventral half of the embryo during the syncytial blastoderm stage of development. NK-2 expression is also regulated through suppression by snail,52 singleminded53-55 and decapentaplegic56,57 in dorsally derived tissues. The mammalian homologues of some of these genes are known, but their role in regulation of Nkx-2.1 requires further studies.

Nkx-2.1 Regulates Lung and Thyroid Pattern Formation The precise role of Nkx 2.1 in lung and thyroid development has been investigated by both in vitro and in vivo techniques.44,58 Suppression of Nkx-2.1 mRNA translation by an antisense oligonucleotide strategy in mouse embryonic lung explants has been shown to abrogate normal branching morphogenesis.58 In these lungs, branches consisted of large baggy structures whose histological examination revealed unorganized dysplastic epithelial cell masses obtruding into the airway space. Consistent with these in vitro observations, the lungs of E18, Nkx-2.1(-/-) mouse embryos are profoundly abnormal and nonfunctional. Phenotypic impact of targeted disruption of the Nkx-2.1 locus is identifiable in lungs of mutant embryos as early as E12. In contrast to wild type lungs, which include several bronchial generations, the Nkx-2.1(-/-) lungs failed to undergo normal branching morphogenesis beyond the mainstem bronchi. At the end of each main-stem bronchus the Nkx-2.1(-/-) dysmorphic lungs consisted of translucent semi-amorphous bags containing what appears to be normal mesenchyme and a multi-layered epithelial lining, formed exclusively of columnar epithelial cells. With the exception of scattered columnar, ciliated epithelial cells, no morphologic and/or biochemical evidence of specific pulmonary cell differentiation could be observed in Nkx-2.1(-/-) lungs. In particular none of the lung epithelial-specific differentiation genes, including surfactant protein genes SP-A, SP-B or SP-C are expressed in Nkx-2.1(-/-) lungs. Thus, lung morphogenesis in Nkx 2.1(-/-) embryos is arrested at the onset of the pseudoglandular period of lung development, resulting in the absence of those distal structures which normally develop subsequent to extensive branching morphogenesis and regional cellular specification. Nkx-2.1 is the first example of a homeodomaincontaining transcriptional protein with a demonstrated role in branching morphogenesis and differentiation of specialized pulmonary epithelial cell types. The role of Nkx-2.1 in thyroid development is clearly seen in the Nkx-2.1(-/-) mouse. These mutant mice completely lack the thyroid.44 The thyroid’s neighboring tissue, the parathyroid, however, which is derived from the third pharyngeal pouches, appears to be normal in these mice. In the wild type mouse embryo, the thyroid primordium is formed at around E9, starts migrating caudally, and completes its bifurcation to give rise to the two lobes of the thyroid gland which flank the trachea by E13.5. This bilateral structure is not formed in E13 Nkx-2.1(-/-) embryos. However, the thyroid primordium is clearly present, although abnormal in appearance in E10-E10.5 mutant embryos. One mechanism that ac-

Endodermal Appendage Formation

209

counts for this phenotype is absence of proliferation of progenitor cells in the Nkx-2.1(-/-) thyroid primordium. Thus, as in the lung, although specification of thyroid primordium occurs in Nkx-2.1(-/-) embryos, thyroid morphogenesis is strictly dependent on the activity of Nkx-2.1. These observations further demonstrate a critical role of Nkx-2.1 in proliferation and differentiation of specialized thyroid epithelial cell types.

Possible Functions for Nkx-2.1 It is now clear that normal morphogenesis of thyroid and lung rudiments is strictly dependent on the wild type activity of Nkx-2.1. Nkx-2.1 encodes a transcriptional factor whose function must ultimately be related to its ability to activate or suppress the expression of downstream genes. One category of such target genes has already been identified by a number of in vitro studies. These are the thyroid and lung differentiation-specific genes. In the thyroid, Nkx-2.1 activates the expression of genes for thyroid peroxidase, thyroglobulin and the receptor for thyroid stimulating hormone.34,35,59-61 In the lung, the genes responsible for the production of three proteins found in the pulmonary surfactant, SP-A, SP-B and SP-C, and the gene encoding the Clara cell secretory protein CC10 are all regulated by Nkx-2.1.28,29,42,43 However, neither the genes targeted by Nkx-2.1 in the thyroid nor those in the lung are known to have morphoregulatory function. Therefore, another class of Nkx-2.1 target genes must exist whose encoded proteins regulate lung and thyroid morphogenesis. The identity of this class remains unknown. However, both in vitro and in vivo results are consistent with the possibility that Nkx-2.1 may be related to the establishment of pattern formation during development of these tissues. What are the possible mechanisms by which Nkx-2.1 could regulate lung and thyroid morphogenesis? In the lung and thyroid, as in other organs, morphogenesis requires spatially and temporally well defined developmental cues. An extensive body of experimental evidence shows that the splanchnic mesenchyme, through epithelial-mesenchymal interactions, plays an instructive role in lung morphogenesis (chapters 10 and 12 in this book include detailed reviews of epithelial-mesenchymal interactions in lung and gut morphogenesis). In this context, one possible function for Nkx-2.1 may be in activation of epithelial cell pathways that are necessary for receiving and or interpreting the instructive signals which originate from the mesenchyme. In this role, downstream target genes for Nkx-2.1 would potentially include those encoding cell-surface receptors, components of the signal transduction pathway and a myriad of other factors connecting the cell surface to changes in gene expression and cellular behavior. In a previous section of this chapter, we discussed the concept of positional information proposed by Wolpert.7,8 Whether instructed by the mesenchyme or originating from the epithelium, developmental cues confer positional information at the level of individual cells to control pattern formation. When assignment or interpretation of positional information is disrupted, cells display inappropriate behavior resulting in abnormal morphogenesis. For example, cells within a tissue may choose to proliferate or migrate when and where their normal positional information would have dictated otherwise. This is exactly as we have observed in Nkx-2.1(-/-) mutant lung and thyroid. In the mutant lungs, epithelial cells which normally form a highly branched structure (the pulmonary tree) instead developed into grossly dilated cysts with no sign of defined branching activity. Similarly, although the progenitor of the thyroid glands formed, these cells failed to undergo morphogenesis in the absence of Nkx-2.1 activity. These data suggest that one possibility is that Nkx-2.1 constitutes a component of a transcriptional regulatory mechanism which is necessary for establishment or interpretation of positional information during lung and thyroid pattern formation and morphogenesis. The future task is to identify the nature of the downstream

210

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 11.1. A simple molecular morphogenetic map for lung and thyroid development.

morphoregulatory target genes through which Nkx-2.1 acts to bring about normal morphogenesis.

A Simple Molecular Morphogenetic Map for Lung and Thyroid Development Based on what has been reviewed above, it is appealing to incorporate the available data into an operational algorithm by which the emergence of the lung and thyroid as foregut endodermal derivatives, and their subsequent morphogenesis, can be molecularly described. In such a scheme, HNF-3α, HNF-3β and HNF-3γ occupy a primary position whose regionally-specific expression along the gut axis divides this structure into anterior and posterior compartments (Fig. 11.1). Although the specific mechanism remains elusive, the lung and the thyroid primordia are specified within the domain demarcated by the expression of HNF-3α and HNF-3β. The second tier of regulation appears to involve members of the Nk family of transcriptional factors.62,63 In particular, expression of Nkx-2.1 occurs specifically in the ventral half of the anterior foregut, thereby further regionalizing the foregut into dorsal and ventral compartments. Although both the lung and the thyroid appendages are derived from the ventral compartment of the anterior foregut, absence of Nkx-2.1 expression in Nkx-2.1 (-/-) embryos does not block their formation. In these embryos, the lungs, although profoundly dysplastic, nevertheless are formed, suggesting that Nkx-2.1 is not required for specification of the lung primordium. Similarly, thyroid rudiments are

Endodermal Appendage Formation

211

formed in Nkx-2.1(-/-) embryos, but are abnormal and subsequently undergo what appears to be apoptosis, further suggesting that specification of thyroid appendages is also not dependent on a wild type Nkx-2.1 allele. It is noteworthy that the other members of the Nk family of transcriptional proteins also appear to function not in determination and specification of the tissue progenitors, but rather in morphogenesis of the tissues in which they are expressed. For example Nkx-2.5 is expressed in the thyroid and the heart.63,64 A homozygous targeted disruption of the Nkx-2.5 locus results, not in the absence of a heart, but in abnormal heart morphogenesis.65 Further distinction between thyroid and the lung is noted by differential expression of other members of the Nkx gene family. Nkx-2.6 expression is reported to be confined to the pharynx and the lung,63,64 whereas expression of Nkx-2.5 occurs in the thyroid and not the lung.63,64 In addition, expression of Pax8, encoding a paireddomain transcriptional regulatory protein which occurs in the thyroid gland and not the lungs, further distinguishes the two tissues from one another.66 Construction of a morphogenetic map for lung and thyroid requires precise and detailed knowledge of individual steps involved in morphogenesis of these tissues. At the time of this review, the number of known steps defined by patterns of transcriptional regulatory gene expression is limited. Nevertheless, mapping of possible hierarchies of morphoregulatory steps based on currently available information will aid in generation of testable hypotheses, thereby stimulating further experimental work in this area (Fig. 11.1). We are hopeful that, because of current interest, additional detailed information concerning the molecular regulatory pathways which govern endodermally-derived tissue morphogenesis will be forthcoming soon, enabling us to fill the gaps in the above proposed map.

Acknowledgment Research in our laboratory is supported by HL56590 and HL56221 from NHLBI, the National Institutes of Health and the Hastings Foundation.

References 1. Curtis D, Apfeld J, Lehmann R. nanos is an evolutionarily conserved organizer of anteriorposterior polarity. Development 1995; 121:1899-1910. 2. Kimble J. An ancient molecular mechanism for establishing embryonic polarity? Science 1994; 266:577-578. 3. Gavis ER, Lehmann R. Translational regulation of nanos by RNA localization. Nature 1994; 369:315-318. 4. Rivera-Pomar R, Kackle H. From gradients to stripes in Drosophila embryogenesis: Filling in the gaps. Trends Genet 1996; 12:478-483. 5. Murata Y, Wharton RP. Binding of pumilio to maternal hunchback mRNA is required for posterior patterning in Drosophila embryos. Cell 1995; 80:747-756. 6. Davidson EH. Later embryogenesis: Regulatory circuitry in morphogenetic fields. Development 1993; 118:665-90. 7. Wolpert L. Positional information and pattern formation. Curr Topics Dev Biol 1971; 6:183-224. 8. Wolpert L. Positional information revisited. Development 1989; Supplement, 3-12. 9. Bienz, M. Homeotic genes and positional signalling in the Drosophila viscera. Trends Genet 1994; 10:22-26. 10. Wood WB, Edgar LG. Patterning in the C. elegans embryo. Trends Genet 1994; 10:49-54. 11. Pederson JD, Kiehart DP, Mahaffey JW. The role of Hom-C genes in segmental transformation: Reexamination of the Drosophila Sex combs reduced embryonic phenotype. Dev Biol 1996; 180:131-142. 12. Warren R, Carroll S. Homeotic genes and diversification of the insect body plan. Curr Opin Genet Dev 1995; 5:459-465.

212

Molecular Basis of Epithelial Appendage Morphogenesis

13. Manak JR, Scott MP. A class act: Conservation of homeodomain protein functions. Development 1994; Supplement 61-77. 14. Reuter R, Scott MP. Expression and function of the homeoeotic genes Antennapedia and Sex combs reduced in the embryonic midgut of Drosophila. Development 1990; 109:289-303. 15. McGinnis W, Krumlauf R. Homeobox genes and axial patterning. Cell 1992; 68:283-302. 16. Dolle P, Izpisua-Belmonte JC, Brown J et al. Hox genes and the morphogensis of the vertebrate limb. Prog Clin Biol Res 1993; 11-20. 17. Krumlauf R. Hox genes in vertebrate development. Cell 1994; 78:191-201. 18. Rugh R. The Mouse, its Reproduction and Development. Minneapolis: Burgess 1968. 19. Hebel R, Stromberg MW. The respiratory system. In: Hebel R, Stromberg MW, eds. Anatomy and Embryology of the Laboratory Rat. Worthsee, Federal Republic of Germany: Biomed Verlag 1986. 20. Ten Have-Opbroek AAW. The development of the lung in mammals: An analysis of concepts and findings. Am J Anat 1981; 162:201-19. 21. Costa RH, Lai E, Grayson DR et al. The cell specific enhancer of the mouse transthyretin gene binds a common factor at one site and a liver-specific factor(s) at two other sites. Mol Cell Biol 1988; 8:81-90. 22. Herbst RS, Nielsch U, Sladek F et al. Differential regulation of hepatocyte enriched transcription factors explains changes in albumin and transthyretin gene expression among hepatoma cells. New Biol 1991; 3:289-296. 23. Jurgens G, Weigel D. Terminal versus segmental development in the Drosophila embryo: The role of the homeotic gene forkhead. Roux’s Arch Dev Biol 1988; 197:345-354. 24. Weigel D, Jurgens G, Kuttner F et al. The homeotic gene forkhead encodes a nuclear protein and is expressed in the terminal regions of the Drosophila embryo. Cell 1989; 57:645-658. 25. Weigel D, Jackle H. The forkhead domain: A novel DNA binding motif of eukaryotic transcription factors. Cell 1990; 63:455-456. 26. Lai E, Prezioso VR, Tao W et al. Hepatocyte nuclear factor 3 belongs to a family in mammals that is homologous to the Drosophila homeotic gene forkhead. Genes Dev 1991; 5:416-27. 27. Monaghan AP, Kaestner KH, Grau E et al. Postimplantation expression patterns indicate a role for the mouse forkhead/HNF-3, and genes in determination of the definitive endoderm, chordamesoderm and neuroectoderm. Development 1993; 119:567-78. 28. Sawaya PL, Stripp BR, Whitsett JA et al. The lung-specific CC10 gene is regulated by transcription factors from the AP-l, octamer, and hepatocyte nuclear factor 3 families. Mol Cell Biol 1993; 13:3860-71. 29. Bohinski RJ, Di Lauro R, Whitsett JA. The lung-specific surfactant protein B gene promoter is a target for thyroid transcription factor 1 and hepatocyte nuclear factor 3, indicating common factors for organ-specific gene expression along the foregut axis. Mol Cell Biol 1994; 14:5671-5681. 30. Sato K, Di Lauro R. Hepatocyte nuclear factor 3 beta participates in the transcriptional regulation of the thyroperoxidase promoter. Biochem Biophys Res Commun 1996; 220:86-93. 31. Ang SL, Rossant J. HNF-3β is essential for node and notochord formation in mouse development. Cell 1994; 78:561-574. 32. Weinstein DC, Ruiz-i-Altaba A, Chen WS et al. The winged-helix transcription factor HNF3β is required for notochord development in the mouse embryo. Cell 1994; 78:575-588. 33. Lazzaro D, Price M, De Felice M et al. The transcription factor TTF-1 is expressed at the onset of thryoid and lung morphogenesis and in restricted regions of the foetal brain. Development 1991; 113:1093-1104. 34. Civitareale D, Lonigro R, Sinclair AJ et al. A thyroid-specific nuclear protein essential for tissue-specific expression of the thyroglobulin promoter. EMBO J 1989; 8:2537-2542. 35. Guazzi S, Price M, De Felice M et al. Thyroid nuclear factor 1 (TTF-1) contains a homeodomain and displays a novel DNA binding specificity. EMBO J 1990; 9:3631-3639.

Endodermal Appendage Formation

213

36. Mizuno K, Gonzalez F, Kimura S. Thyroid-specific enhancer-binding protein (T/ebp): cDNA cloning, functional characterization, and structural identity with thyroid transcription factor TTF-1. Mol Cell Biol 1991; 11:4927-4933. 37. Endo T, Ohta K, Saito T et al. Structure of the rat thyroid transcription factor-1 (TTF-1)gene. Biochem Biophys ResCommun 1994; 204:1358-1363. 38. Oguchi H, Pan YT, Kimura S. The complete nucleotide sequence of the mouse thyroidspecific enhancer-binding protein (T/ebp) gene: Extensive identity of the deduced amino acid sequence with the human protein. Biochim Biophys Acta 1995; 1261:304-306. 39. Ikeda K, Clark JC, Shaw-White JR et al. Gene structure and expression of human thyroid transcription factor-1 in respiratory epithelial cells. J Biol Chem 1995; 270:8108-8114. 40. Kim Y, Nirenberg M. Drosophila NK-homeobox genes. Proc Natl Acad Sci USA 1989; 86:7716-7720. 41. Damante G, Fabbro D, Pellizzari L et al. Sequence-specific DNA recognition by the thyroid transcription factor-l homeodomain. Nuc Acids Res 1994; 22:3075-3083. 42. Kelly SE, Bachurski CJ, Burhans MS et al. Transcription of lung-specific surfactant protein C gene is mediated by thyroid transcription factor-1. J Biol Chem 1996; 271:6881-6888. 43. Bruno MD, Bohinski RJ, Huelsman KM et al. Lung cell-specific expression of the murine surfactant protein A (SP-A) gene is mediated by interactions between the SP-A promoter and thyroid transcription factor-1. J Biol Chem 1995; 270:6531-6536. 44. Kimura S, Hara Y, Pineau T et al. The T/ebp null mouse: thyroid-specific enhancer-binding protein is essential for the organogenesis of the thyroid, lung, ventral forebrain and pituitary. Genes Dev 1996; 10:60-69. 45. Skandalakis JE, Gray SW, Ricketts R. The esophagus. In: Skandalakis JE, Gray SW, eds. Embryology for Surgeons. Baltimore: Williams and Wilkins, 2nd edition. 1994. 46. Minoo P, King RJ. Epithelial-mesenchymal interactions in lung development. Annu Rev Physiol 1994; 56:13-45. 47. Minoo P, Liu H, Bringas P et al. Nkx-2.1 is required for dorso-ventral patterning of the anterior foregut. (Submitted). 48. Mellerick DM, Nirenberg M. Dorsal-ventral patterning genes restrict NK-2 homeobox gene expression to the ventral half of the central nervous system of Drosophila embryos. Dev Biol 1995; 171:306-316. 49. Roth S, Stein D, Nusslein-Volhard C. A gradient of nuclear localization of the dorsal protein determines dorsoventral pattern in Drosophila embryos. Cell 1989; 59:1189-1202. 50. Rushlow CA, Han K, Manley JL et al. The graded distribution of the dorsal morphogen is initiated by selective nuclear transport in Drosophila. Cell 1989; 59:1165-1177. 51. Steward R, Govind S. Dorsal-ventral polarity in the Drosophila embryo. Curr Opin Genet Dev 1993; 3:556-561. 52. Boulay JL, Dennefeld C, Alberga A. The Drosophila developmental gene snail encodes a protein with nucleic acid binding fingers. Nature 1987; 330:395-398. 53. Mayer U, Nusslein-Volhard C. A group of genes required for pattern formation in the ventral ectoderm of Drosophila embryo. Genes Dev 1988; 2:1496-1511. 54. Thomas JB, Crews ST, Goodman CS. Molecular genetics of the single-minded locus: A gene invovled in the development of the Drosophila nervous system. Cell 1988; 52:133-141. 55. Crews ST, Thomas JB, Goodman CS. The Drosophila single-minded gene encodes a nuclear protein with sequence similarity to the per gene product. Cell 1988; 52:143-151. 56. Spencer FA, Hoffman FM, Gelbart WM. decapentaplegic: A gene complex affecting morphogenesis in Drosophila melanogaster. Cell 1982; 28:451-461. 57. Padgett RW, St. Johnston RD, Gelbart WM. A transcript from a Drosophila pattern gene predicts a protein homologous to the transforming growth factor β family. Nature 1987; 325:81-84. 58. Minoo P, Hamdan H, Bu D et al. TTF-1 regulates lung epithelial morphogenesis. Dev Biol 1995; 172:694-698. 59. Zannini M, Francis-Lang H, Plachov D et al. Pax8, a paired domain-conating protein binds to a sequence overlapping the recognition site of a homeodomain and activates transcription from two thyroid-specific promoters. Mol Cell Biol 1992; 12:4230-4241.

214

Molecular Basis of Epithelial Appendage Morphogenesis

60. Francis-Lang H, Zannini M, de felice M et al. Multiple mechanisms of interference between transformation and differentiation in thyroid cells. Mol Cell Biol 1992; 12:5793-5800. 61. Shimura H, Okajima F, Ikuyama S et al. Thyroid-specific expression and adenosine 3', 5'-monophosphate autoregulation of the thyrotropin receptor gene involves thyroid transcription factor-1. Mol Endocrinol 1994; 8:1049-1069. 62. Burglin TR. A comprehensive classification of homeobox genes. In: Guidebook to the Homeobox Genes. Duboule D, ed. Oxford: Oxford Univ press, 1993: 25-71. 63. Harvey RP. NK-2 homeobox genes and heart development. Dev Biol 1996; 178:203-216. 64. Lints TJ, Parsons LM, Hartley L et al. Nkx-2.5: A novel murine homeobox gene expressed in early heart progenitor cells and their myogenic descendants. Development 1993; 119:419-431. 65. Lyons I, Parsons LM, Hartley L et al. Myogenic and morphogenetic defects in the heart tubes of murine embryos lacking the homeobox gene Nkx-2.5. Genes Dev 1995; 9:1654-1666. 66. Plachov DK, Chowdhury K, Walther C et al. Pax8, a murine paired box gene expressed in the developing excretory system and thyroid gland. Development 1990; 110:643-651.

CHAPTER 12

Lung Epithelial Morphogenesis: Integrated Functions of Transcriptional Factors, Peptide Growth Factors, Extracellular Matrix, Physiological and Environmental Factors David Warburton, Guillermo Flores-Delgado, Ding Bu, Kathryn D. Anderson and Richard E. Olver “A remarkable engineering feat provides a vast and extremely thin surface for the transfer of gases between air and blood.....how little we know ‘for sure’ and how much remains to be learned.” Julius H. Comroe, Jr., 1965 “In the new field of Developmental Biology, things will seem more complicated before the simplicities become apparent.” Wilhelm Roux, 1896.

Introduction

T

he mammalian lung has the largest surface area of any epithelial appendage in the human body. Its intricate vaulted structure could indeed be considered a veritable “biological cathedral”. The lung develops as an epithelial appendage of the primitive foregut in the early embryo, undergoes extensive branching morphogenesis in utero and continues to undergo alveolarization and increases in surface area postnatally. The human lung achieves a final gas diffusion surface area of 70 m2 by 0.1 µm thick in young adulthood. This is an area about 40 times the surface area of the next largest epidermal appendage, the skin. It is capable of supporting a systemic oxygen consumption ranging between 250 ml/min at rest to 5500 ml/min during exercise. A matching capillary network also develops in close apposition to the alveolar surface which can accommodate a blood flow rising from 4 to 40 L/min during the transition from rest to maximal exercise.1 The lung originates as a ventral appendage of the endodermal epithelium lining the floor of the primitive embryonic anterior pharynx at E9.5 days in the mouse (Fig. 12.1). It then divides laterally into two buds and begins dichotomous branching into the surrounding splanchnic mesenchyme. This repetitive epithelial branching process, termed branching morphogenesis, is characteristic of lung formation and continues throughout gestation. Histologically, lung development has been divided into four chronological stages in the mouse:

Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

216

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 12.1. Simplified diagram illustrating the major events in murine lung epithelial morphogenesis The primitive lung epithelial bud forms from the ventral surface of the embryonic foregut at E9.5; the lobes and major bronchi have formed by E11; dichotomous branching of the airways continues between E11-16; saccules form at the ends of the airways and are partitioned into alveoli between E16 and postnatal day 30.

1. pseudoglandular stage (E9.5-E16.6), during which the bronchial and respiratory tree develops and an undifferentiated primordial system forms; 2. canalicular (E16.6-E17.4)—terminal sacs and vascularization develop in this period; 3. terminal sac stage (E17.4-postnatal day 5 (P5))—number of terminal sacs and vascularization increase and type I and II cells differentiate; and 4. alveolar stage (P5-P30)—terminal sacs develop into mature alveolar ducts and alveoli. Among these stages, the most significant growth and branching of the primitive lung epithelium take place in the pseudoglandular stage, during which the initial pattern of the lung and the complete respiratory tract forms. Thus, the branching pattern and organization of lung epithelial cells differ between embryonic and fetal lung development. Embryonic lung branching is dichotomous and establishes the bronchial airway tree, while fetal branching morphogenesis results in the formation of sacs and alveoli along the distal-most branches of the bronchial tree (Fig. 12.1). The lung is a relatively ancient epithelial appendage to appear in evolution: Lungs facilitated the transition from aquatic to terrestrial existence. The lungs confer an evolutionary advantage by presenting an extensive, thin surface area, which facilitates gas exchange between the atmosphere and the perfusate of the organism. Alternative evolutionary strat-

Lung Epithelial Morphogenesis

217

egies to solve the problem of gas diffusion to support tissue respiration include: direct diffusion through the body surface in very small organisms; tubules termed tracheae which invaginate from the lateral surface of the body in insects such as Drosophila larva; and diffusion of gases in solution from the aqueous environment to the organismal perfusate, which occurs through the gills in fish. The lungs of small air breathing creatures such as the lung fish and frog are primitive sac-like structures which are not extensively branched, yet present sufficient surface area to support their relatively low rate of oxygen consumption. Larger homeothermic air breathing creatures such as the Mammalia have perforce developed much more extensively branched lungs, which present a very large surface area for gas diffusion. In order to solve the developmental problem of forming an organ capable of conducting respiratory gases to a large, diffusible interface with the circulation, the lung epithelium must undergo cell proliferation to increase its surface area, branching morphogenesis and alveolar saccule formation, as well as cell lineage differentiation. These developmental processes are determined by well-coordinated epithelial-mesenchymal interactions between transcriptional factor mediated mechanisms, peptide growth factor signaling, cell cycle control mechanisms and extracellular matrix expression and signaling (Fig. 12.2). The goal of this chapter is to review what is currently known about the molecular basis for the epithelial-mesenchymal interactions that mediate the induction of morphogenesis in the embryonic mouse and, by inference, the human lung. Herein, we focus on the role of specific transcriptional factors, peptide growth factor signaling mechanisms, extracellular matrix components and cell adhesion molecules that play key instructive and permissive roles. The impact of physiological and environmental factors is also integrated into the discussion of lung morphogenesis. We then suggest and discuss how novel research directions which are already proving fruitful to further elucidate the molecular basis of lung morphogenesis may lead to new rational therapeutic approaches to inducing lung regeneration in human disease states such as infantile chronic lung disease, pulmonary hypoplasia and chronic emphysema.

Peripheral Embryonic Lung Mesenchyme as a “Compleat” Inducer of Lung Morphogenesis The linked concepts of morphogens and morphogenetic gradients within multicellular developing organisms have been extant for a century.2 The currently accepted hypothesis that morphogenetic signals instruct differential gene expression between cell types during embryonic morphogenesis originates in the classical observation of the “Spemann organizer” in the dorsal lip of the blastopore in Xenopus embryos.3 This paradigm has now been demonstrated to be generally true in the organogenesis of such diverse systems as the limb bud, tooth, breast, pancreas, kidney, gut, neural tube and heart. A recent refinement of these concepts demonstrates that specific morphogens can act both at short and long range within a morphogenetic gradient, depending on whether the morphogen is tethered or diffusible.4 The early embryonic lung develops at E9.5 in the mouse as an endodermal epithelial appendage which arises in the midline from the ventral surface of the primitive foregut and extends caudally and ventrally into the surrounding primitive mesenchyme. Rostro-caudally, these structures form the primitive larynx, trachea and mainstem and lobar bronchi by E10-11 in the mouse. Subsequent branching morphogenetic events give rise to the segmental and subsegmental generations of bronchi, respiratory bronchioles, terminal bronchioles and finally the alveoli in the left and right lungs. Both classical and more recent mesenchyme recombination experiments demonstrate that the induction of early embryonic lung branching morphogenesis is determined by soluble factors produced by the primitive peripheral lung mesenchyme. Transplantation of peripheral lung mesenchyme to the E11 mouse trachea in culture results not only in the

218

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 12.2. Epithelial-Mesenchymal interactions in lung epithelial morphogenesis. To solve the developmental problem of forming an organ capable of conducting respiratory gases to a large, diffusible interface with the circulation, the lung epithelium must undergo cell proliferation to increase its surface area, branching morphogenesis and alveolar saccule formation, as well as cell lineage differentiation. These developmental processes are determined by well-coordinated epithelial-mesenchymal interactions between transcriptional factor mediated mechanisms, peptide growth factor signaling, cell cycle control mechanisms and extracellular matrix expression and signaling. Physiologic and environmental factors also regulate these processes..

induction of supernumerary pulmonary branches from the trachea, but also induction of genes which mark the expression of peripheral epithelial cell lineages5-7 (Fig. 12.3). A number of soluble factors have been adduced as potential mediators of the morphogenetic effects of mesenchyme upon the primitive lung epithelium, including EGF, IGF, bFGF, KGF/FGF-7, HGF and PDGF.8-10 The extracellular matrix also plays a key role in determining branching morphogenesis of the lung, and it has been suggested that insoluble macromolecules of the basal lamina and deeper matrix form physical barriers or traps to sequester soluble components, which can in turn regulate epithelial and mesenchymal cell proliferation.11

Lung Epithelial Morphogenesis

219

I TTF-1? SP-A SP-C

MESENCHYME RECOMBINATION II A

B

Fig. 12.3. Transplantation of peripheral lung mesenchyme to the E11 mouse trachea in culture results in the induction of supernumerary pulmonary branches and expression of peripheral epithelial specific genes in these supernumerary branches of epithelium. This is illustrated diagrammatically in panel I. In panel II, an E11 mouse lung is shown with peripheral mesenchyme removed, exposing primitive epithelium (arrows) and tracheal mesenchyme removed prior to the transplantation of the peripheral mesenchyme to the proximal trachea (arrow heads). The same embryonic lung explant is shown in B after 4 days in culture. The arrow heads indicate where two supernumerary epithelial branch systems have been induced to arise from the proximal trachea. In contrast, the exposed peripheral mesenchyme has failed to branch any further (arrow heads).

Genetic and Epigenetic Factors Including Key Transcriptional Factors, Peptide Growth Factor Receptor-Mediated Signaling Pathways, Extracellular Matrix, Integrins and Environmental Influences Regulate Lung Morphogenesis These inputs are integrated during the normal process of embryonic, fetal and postnatal lung organogenesis to determine organized patterns of cellular proliferation and cell lineage differentiation which eventually correlate structure with physiological function. Thus, lung development extends in a coordinated manner from branching morphogenesis in early embryonic life, through the critical transition from fetal life to air breathing, up to the completion of alveolarization which occurs postnatally. Pulmonary branching is reproducible as far down the respiratory tree as the sub-segmental bronchi. Between that level and the alveoli, branching appears to be governed by fractal functions. Alveoli are formed by epithelial sheets folding at the sites of expression of elastin fibers. The process of morphogenesis can also be positively or negatively impacted by physiological and environmental factors such as lung fluid volume, mechanical strain and hyperoxia (Fig. 12.2).

220

Molecular Basis of Epithelial Appendage Morphogenesis

Lessons on Branching Morphogenesis from Drosophila Genetics Branching Mutants of the Drosophila Respiratory Tracheae May Be Functional Paradigms for Branching Morphogenesis in General Drosophila embryos solve the problem of tissue respiration by delivering air directly to individual tissues through a series of tubes termed tracheae. The tracheae originate during Drosophila embryogenesis from the respiratory placodes which are placed segmentally along the lateral surface of the embryo. The placodes cells proliferate to a limited extent and invaginate inwards to form initial branches. Cytoplasmic projections of individual tracheal cells then ramify to form a series of gas delivery tubules which terminate in a predictable fashion at individual internal organs. At least 30 tracheal enhancer trap markers and several distinct mutations in Drosophila are associated with specific defects in a sequential series of morphologically distinct but genetically coupled branching events in the process of tracheal branching morphogenesis.12 Recently, trachealess (trh) has been identified as necessary to direct the initial steps of tubulogenesis in both the respiratory organs and salivary glands of Drosophila.13 In trh mutants, tube-forming cells of the trachea, salivary gland and filzkörper fail to invaginate to form tubes and remain on the embryo surface.14 trh expression is in turn controlled by the Sex combs reduced (scr) and forkhead (fkh) transcriptional factors, and is homologous to the human hypoxia-inducible factor-1.15 Branchless (bnl) encodes a Drosophila FGF ligand homologue that controls tracheal cell migration and hence the pattern of branching.16 The latter mutation is complimentary to the breathless (brl) mutation, which is a loss of function mutation in the Drosophila brl receptor, which is homologous to the mammalian FGFR. Interestingly, expression of a dominant negative FGFR driven by the SP-C promoter in transgenic mice also results in severe lung hypoplasia, comprising loss of bronchial branching distal to the mainstem bronchi and of pulmonary angiogenesis.17 The phenotype of the bnl and brl loss of function mutations suggest that the FGF pathway directs the migration of tracheal cells during primary branching and then activates downstream programs of finer branching at the ends of the growing branches.16 In the pointed loss of function Drosophila mutant pnt∆88, primary tracheal branching occurs normally, but no secondary branches are ever formed. The pnt gene is also required for activating and repressing marker genes that underly terminal branching and branch fusion. The pnt gene encodes two ETS-like proteins which are also involved in the development of midline glial cells.18 The pruned gene product is a Drosophila homologue of serum response factor (SRF), which functions as a growth factor-activated transcription complex together with an ETS domain ternary complex factor.19 In pruned loss of function mutants, terminal tracheal cells failed to extend cytoplasmic projections, while with expression of either a constitutively activated SRF or ternary complex factor, extra projections grow out in an unregulated manner.19 Thus, Drosophila SRF may function in a growth factor inducible transcription complex to regulate distal cytoplasmic outgrowths during tracheal epithelial branching morphogenesis. Branching of the respiratory tracheae in Drosophila can be considered as a paradigm for morphogenesis of branched epithelial appendages in general. It will be interesting to determine whether mammalian homologues of these additional Drosophila gene families also play a role in diseases affecting other branching epithelial appendages in humans, in addition to the lung.

Lung Epithelial Morphogenesis

221

Fig. 12.4. A flow diagram depicting the histological classification of stages in lung morphogenesis, with known molecular determinants of lung morphogenesis superimposed at the approximate stage at which each null mutation (-/-), hSP-C promoter driven transgene, exogenous growth factor, dominant negative receptor, antisense oligonucleotide or antibody is thought to exert its effect. For a more detailed explanation of the role of each of these molecular determinants in lung morphogenesis, please consult the relevant section of the text. FGFR∆K- is a dominant negative FGFR with deletion in the tyrosine kinase domain. SP-C- indicates a transgene driven by the human surfactant protein-C promoter. TGF-β IIR AS, Ab indicates abrogation of transforming growth factor type II function by antisense oligonucleotides or by immunoperturbation with blocking antibodies.

Molecular Determinants of Lung Morphogenesis Known molecular determinants of lung morphogenesis are summarized in Figure 12.4. In this figure are presented the currently published null mutations (-/-), SP-C transgenic mouse lines, and exogenous growth factors which are known to exert effects on lung morphogenesis. The respective roles of these individual factors is discussed below.

Transcriptional Factors Termed Master Genes Provide the Developmental Genetic Instructions for Pulmonary Organogenesis The pattern forming events that control the initial and subsequent steps of pulmonary organogenesis are only now beginning to be understood. We and others have recently demonstrated that the thyroid transcription factor-1 (TTF-1) family, also called the thyroid enhancer binding protein (T/ebp) transcriptional factor family and Nkx-2.1, is essential for the complete induction of embryonic lung branching morphogenesis.20,21 Abrogation of TTF-1 expression with antisense oligodeoxynucleotides results in complete interruption of branching morphogenesis and dysplasia of the embryonic mouse lung epithelium in culture.20 Null mutation of TTF-1 results in a very similar neonatal lethal lung phenotype in association with absence of the thyroid, pituitary and parts of the brain.21 Recently, Minoo and coworkers have noticed that TTF-1 (Nkx-2.1) null mutation also results in failure of septation of the esophago-tracheal primordium, resulting in a phenotype resembling severe human cases of complete tracheo-esophageal fistula (please see chapter 11 for further details of the Nkx-2.1 null mutant phenotype). TTF-1 consensus recognition sites are found in the 5' promoters of several peripheral lung cell lineage specific genes, including SP-A, SP-B, SP-C, SP-D, CC10 and TTF-1 itself.22-27

222

Molecular Basis of Epithelial Appendage Morphogenesis

However, while TTF-1 expression appears to be necessary for lung morphogenesis, other families of transcription factors are also clearly involved. The hepatocyte nuclear (HNF) family of transcription factors are related to the forkhead family of Drosophila and are known to play key roles in regional specification of epithelial cell fates in the gastrointestinal tract and liver. HNF-α is expressed in the gut epithelium anterior to the liver, and consensus HNF binding sites are found in the 5' promoter regions of peripheral lung specific genes, including SP-A, -B, -C, -D and CC10, in close proximity to TTF-1 sites.23,28 Thus, the HNF family appears to cooperate with the TTF-1 family to determine pulmonary epithelial cell lineage fates.29,30 The role of transcriptional factors in the formation of appendages of the early anterior pharynx is considered in greater detail in chapter 11. However, how these morphogenetic transcriptional factors regulate and are regulated by peptide growth factor signaling mechanisms is largely unknown. Recently we have found that TTF-1 is positively regulated by EGFR signaling, as is pulmonary morphogenesis per se.

Peptide Growth Factor Signaling Is Both Inductive and Permissive for Lung Morphogenesis and Modulates Transcriptional Mechanisms Branching morphogenesis and cell lineage differentiation occur spontaneously in E11 mouse early embryonic lung under serumless, chemically defined conditions as well as in zero gravity,9,31-33 suggesting the hypothesis that autocrine and paracrine factors produced within the lung anlage are necessary and sufficient for at least the embryonic phase of lung branching morphogenesis to occur.9,31-33 Soluble factors released by peripheral lung mesenchyme can induce ectopic branching from the trachea of early mouse embryonic lung explants, as well as inducing expression of a complete repertoire of genes specific to peripheral lung epithelium, including SP-A,-B, -C and CC10.7 Candidate inductive and permissive peptide growth factors whose signaling peptides and cognate receptors are expressed in the early mouse embryonic lung include EGF, bFGF, HGF, IGF, KGF, PDGF and TGF-β3. Their inductive and/or permissive influences on lung development have been demonstrated by gain and loss of function experiments in early embryonic mouse lung organ culture, and in transgenic and in null mutant mice.9,10,17,34-40 In general, peptide growth factor cognate receptors with tyrosine kinase intracellular signaling domains such as EGFR, FGFR, c-Met, IGFR, KGFR and PDGFR stimulate lung morphogenesis, while those cognate receptors with serine/threonine kinase intracellular signaling domains, such as the TGF-β family are inhibitory.9,36,37,41-45 The functional evidence supporting the involvement of each of these classes of peptide growth factors and their cognate receptors will be considered in turn. EGF, TGF-α, amphiregulin and the EGF receptor EGF and TGF-α and amphiregulin are all EGFR ligands which can positively modulate early mouse embryonic lung branching morphogenesis.9,41,46 Stimulation of EGFR signaling with exogenous ligand stimulates early murine embryonic lung branching morphogenesis, 9,46 while abrogation of EGFR signaling with tyrphostins, EGFR antisense oligodeoxynucleotides, artificially induced maternal immunity or EGFR null mutation all result in decreased branching morphogenesis in culture and a neonatal pulmonary lethal phenotype in the null mutant.9,41,37,47,48 EGF also is expressed in mature alveolar epithelial cells and regulates type 2 cell proliferation through an apparent autocrine mechanism both in culture and in vivo.34 However, respiratory epithelial cell expression of TGF-α using the SP-C promoter in transgenic mice induces postnatal lung fibrosis (Korfhagen et al, 1994).49 We have recently found that branching morphogenesis is reduced by 50% both in vivo and

Lung Epithelial Morphogenesis

223

in vitro in the lungs of EGFR null mutant mice, while addition of exogenous EGF stimulates branching 3-fold, and SP-C levels 50-fold, in wild type embryonic lungs in culture, whereas the EGFR null mutants do not respond at all to exogenous EGF. KGF, FGFs and the FGF receptors As discussed above with regard to the Drosophila bnl and brl mutations, expression of a truncated, dominant negative, kinase deficient FGFR mutant in the primitive respiratory epithelium of transgenic mice results in a lethal phenotype comprising complete pulmonary aplasia distal to the mainstem bronchi.17 Thus, FGFR signaling is clearly essential for distal pulmonary epithelial branching morphogenesis and differentiation, as well as embryonic pulmonary angiogenesis. On the other hand, over-expression of KGF/FGF-7 from the same promoter in transgenic mice results in pulmonary malformations resembling cystic adenomatoid malformation.50 KGF/FGF-7 treatment of murine lung cultures results in a similar phenotype.51,50 The dilated, cystic phenotype appears to be related to decreased expression of α-ENAC, and hence an increase in non-CFTR-dependent fluid secretion in response to KGF.52 On the other hand, acidic FGF can support branching of primitive pulmonary epithelium in defined culture in the absence of mesenchyme.53,54 Intratracheal KGF/FGF-7 also stimulates alveolar epithelial cell proliferation and protects the lung against oxidant injury postnatally.55 Thus, activation of FGFR by FGF family peptides is a prime candidate for a major physiologic role in mediating the inductive effects of mesenchyme on the primitive pulmonary epithelium. HGF and c-Met While HGF/scatter factor was originally isolated as a potent mitogen for hepatocytes, it also has potent motogenic, mitogenic and morphogenetic activities on epithelial cells, including respiratory epithelial cells in vitro.56,57 HGF is expressed in primitive lung mesenchyme, while its receptor, the c-Met tyrosine kinase is expressed in primitive lung epithelium, suggesting the possibility of inductive mesenchymal-epithelial interactions. Expression of HGF in Lx-1 lung cancer cells induces alveolar differentiation.58 The IGF peptides, binding proteins and receptors The insulin-like growth factors (IGFs), their binding proteins and receptors are expressed in both rodent and human fetal lung.59-63 Because there is little apparent change in the distribution or abundance of IGF-I and -2 and their receptors during gestation, but the IGFbps 1-6 are differentially regulated, it is possible that the latter may play a key role in mediating temporo-spatial IGF signaling, particularly the regulation of rates of cellular proliferation.61 Mice carrying null mutation of the insulin-like growth factor-1 (Igf-1) gene are dwarfed to 60% of normal birth weight.64 Depending on the genetic background, some of the Igf-1 null mutants die in the neonatal period, while others survive to reach adulthood. On the other hand, null mutants for the cognate type 1 insulin-like growth factor receptor (Igf1r) gene always die at birth of respiratory failure and exhibit a more severe growth deficiency (45% of normal birth weight). Dwarfism is further exacerbated (30% of normal size) in Igf-1 and Igf-2 double null mutants and in Igf1r and Igf-2 double null mutants. There does not appear to be a gross defect in primary branching morphogenesis per se; the lungs merely appear hypoplastic.64 However, it also seems likely that IGF signaling may play a key role in facilitating signaling by other peptide growth factor pathways involved in lung morphogenesis, since IGF-IR signaling function is required for both the mitogenic and transforming activities of the EGF receptor.65

224

Molecular Basis of Epithelial Appendage Morphogenesis

PDGF peptide and cognate receptor isoforms PDGF peptides are dimeric ligands formed from two peptide chains—A and B. The PDGF-AA and PDGF-BB homodimers and PDGFR are present in embryonic mouse lung and are differentially regulated in fetal rat lung epithelial cells and fibroblasts.66 PDGF-AA regulates both DNA synthesis and early branching in early mouse embryonic lung epithelium in culture: abrogation of PDGF-A expression with antisense oligodeoxynucleotides or PDGF-A blocking antibodies each decreases DNA synthesis, and hence the size of early embryonic mouse lung in culture, as well as interfering with early branch point formation.39 On the other hand, abrogation of PDGF-B chain expression with antisense oligodeoxynucleotides reduces the size of the epithelial component of early embryonic mouse lung explants, but does not reduce the number of branches. PDGF-A homozygous null mutant mice are lethal either prenatally before E.10, or postnatally. Surviving PDGF-A -/- mice develop pulmonary emphysema secondary to failure to septate alveoli.67 This phenotype is apparently caused by loss of alveolar myofibroblasts and associated elastin fiber deposition. Since PDGF-α receptors are expressed in the lung at the location of putative alveolar myofibroblasts, and the latter were specifically absent in PDGF-A null mutants, it appears that PDGF-A chain expression is essential for the ontogeny of pulmonary alveolar myofibroblasts. Thus, PDGF signaling appears to play a permissive role for epithelial DNA synthesis during embryonic life and an instructive role for the ontogeny of myofibroblasts, elastin synthesis by the latter cell lineage and hence alveolarization in postnatal life. It is also interesting to note that PDGF-B chain expression is also essential for the ontogeny of renal mesangial cells. Thus, myofibroblastic cells appear to have key morphogenetic functions in the formation of tubular epithelial appendages both in the lungs and the kidney. Bombesin-like peptides and GRP-receptors GRP is the mammalian equivalent of Bombesin and is produced by pulmonary neuroendocrine (PNE) cells in the adult lung. Expression of GRP peaks during fetal life in the rapidly proliferative phase of airway epithelial development. At earlier stages of development in the mouse, GRP is expressed in undifferentiated epithelial lineage precursor cells.32 Blockade of GRP action by immunoperturbation or pharmacological blockers both in vivo and in culture retards lung epithelial development.68 Thus, GRP-receptor signaling may play a physiological role in the induction of lung morphogenesis, particularly at later fetal stages when PNE cells have differentiated. TGF-β family peptides and cognate receptors TGF-β 1, 2 and 3 peptides and the TGF-β type I and type II receptors are expressed and differentially distributed in the embryonic and fetal lung.69,44,45 TGF-β1 and TGF-β2 both inhibit pulmonary branching morphogenesis in culture, although TGF-β2 is considerably more potent than TGF-β1.42,45 The expression of pRb is not necessary for the inhibitory effects of TGF-β on branching to be transduced, since TGF-β1 inhibits branching in pRb -/embryonic lungs to the same extent as in wild type. However, N-myc expression is suppressed by TGF-β1 in wild type lungs, but is not suppressed by TGF-β in the pRb null mutant, indicating that TGF-β is necessary for the inhibitory effect on n-myc expression to occur.43 Interestingly, gene targeting of the n-myc locus has produced a hypoplastic, neonatal lethal lung phenotype, as well as a lethal hypoplasia of sub-endocardial myoblasts.70 Perhaps the n-myc knockout prevents the pulmonary epithelium from expanding to a sufficient surface area to support gas diffusion postnatally. TGF-β3 null mutation also results in a specific immature-appearing neonatal lung phenotype which is rapidly fatal in newborn mice.36 Unlike the normal neonatal lung pheno-

Lung Epithelial Morphogenesis

225

type found in TGF-β1 null mutant mice, which has been attributed to maternal transplacental rescue, TGF-β3 null mutation appears to be refractory to maternal transplacental rescue. TGF-β3 gene expression is also strongly induced in response to corticosteroid treatment of fetal lung fibroblasts,71 suggesting the hypothesis that the well recognized maturation enhancing effects of glucocorticoids on late fetal lung may in part be mediated by stimulation of temporo-spatially restricted TGF-β3 gene expression. Evidence from normal and targeted misexpression studies in mice further suggests that BMP-4, another TGF-β family peptide, also plays a role in embryonic lung morphogenesis.72 Misexpression of Bmp-4 driven by the SP-C promoter in transgenic mice results in lungs that are smaller than normal, with grossly distended terminal buds and large air-filled sacs at birth. There is also reduced expression of alveolar epithelial cell differentiation markers (SP-C), but not of Clara cell markers (CC10). However, targeted misexpression of TGF-β1 using the same SP-C promoter system also results in a neonatal lethal, hypoplastic pulmonary phenotype with decreased saccule formation and epithelial differentiation.52 Taken together, these findings suggest that TGF-β family peptide over-expression in vivo merely reflects a default negative regulatory effect on morphogenesis similar to that elicited in culture.42,43,45 On the other hand, abrogation of TGF-β type II receptor signaling, either with antisense oligodeoxynucleotides or with blocking antibodies, stimulates lung morphogenesis 2- to 3-fold and increases expression of TTF-1 and SP-C.45 Thus, endogenous autocrine/paracrine TGF-β signaling through the TGF-β type II receptor appears to negatively regulate lung organogenesis. The negative effect of TGF-β signaling through the TGF-β type II receptor on cell cycle progression in pulmonary epithelial cells probably plays a major role in the latter inhibitory effect by limiting expansion of the epithelial surface area. Recent experiments from our laboratory indicate that TGF-β signaling through the TGF-β type I receptor specifically instructs the formation of branch points. Abrogation of TGF-β type I receptor expression with antisense oligodeoxynucleotides significantly reduces the formation of new branch points by E11 early embryonic mouse lung in culture, resulting in a phenotype characterized by long, tubular appearing airways devoid of new branch points. This effect is associated with decreased fibronectin and matrix Gla protein gene expression and failure to form condensations of extracellular matrix containing fibronectin, as would be expected to occur at sites where a new branch point is about to form.73 Moreover, the growing points of the tubular-appearing airways are virtually devoid of fibronectin. Thus, the regulation of fibronectin gene expression by the TGF-β type I receptor apparently plays a key role in the molecular basis of lung morphogenesis by instructing the formation of new airway branch points and the localized deposition of extracellular matrix components. The downstream TGF-β receptor signaling pathway is not as yet completely clear. However, several molecules have been found to impart TGF-β signals. Studies have shown that TGF-β effects on DNA synthesis are pertussis-toxin sensitive, implicating G-proteins as signal transducers.74,75 Immunophilin FKBP-12, a target of the macrolides FK506 and rapamycin, was found to interact with type I receptors of TGF-β family.76 A WD-domain protein, TRIP-1 (TGF-β-receptor interacting protein-1), was found to associate specifically with type II TGF-β receptor in a kinase-dependent way, which may be analogous to the function of GRB-2 of receptor tyrosine kinases.77 Recently, a mouse protein kinase of the MAPKKK family, TAK1 (TGF-β-activated kinase 1), was found to be specifically activated by members of the TGF-β superfamily of ligands and involved in regulation of transcription by TGF-β.78 More recently, a group of genes homologous to Drosophila Mad (mothers against dpp), a gene that interacts with the TGF-β superfamily member encoded by decapentaplegic (dpp), has been found to be cytoplasmic effectors for the TGF-β like signals.79-82 Among them the mammalian homologues Smad3 and Smad4 are thought to be

226

Molecular Basis of Epithelial Appendage Morphogenesis

effectors of TGF-β receptors, in which a Smad3 dimer is associated with a TGF-β receptor complex, and can be phosphorylated by the receptor kinases and then bind with Smad4 which then is translocated to the nucleus to act as a tumor suppressing transcription factor (Zhang et al, 1996).83 It will be interesting to determine the role of the TGF-β pathway signal transduction molecules termed Smads which include transcriptional activators downstream of the TGF-β receptors. In addition, TGF-β interacting proteins and proteoglycans such as betaglycan (TGF-β type III receptor), biglycan, decorin, fibromodulin and endoglyn may affect pulmonary morphogenesis.84 As an examplar, endoglyn is a dimeric TGF-β1 and -3 binding protein of endothelial cells, modulates cellular responses to TGF-β1 and can form heteromeric complexes with TGF- β signaling receptors. 85,86 Interestingly, endoglyn is the gene for Osler-Weber-Rendu hereditary telangiectasia type 1, a condition which is characterized by large intrapulmonary arteriovenous malformations.87 It is also interesting to note that massively dilated pulmonary vessels with thin or absent smooth muscle layers were a prominent pathologic feature of the neonatal lethal TGF-β3 null mutant phenotype.36 Sonic hedgehog (Shh) and patched (ptc) Sonic hedgehog is the murine homologue of the Drosophila hedgehog gene which encodes a secreted protein that regulates Decapentaplegic (Dpp) and wingless (Wg) in target cells.88-91 Low levels of Shh are expressed throughout the primitive respiratory epithelium, with high levels seen at the tips.92,72,93 Patched (ptc) is the murine homologue of Patched (Ptc), a segment polarity gene in Drosophila, which encodes a putative multiple pass membrane spanning receptor which is required for Shh signaling. Ectopic expression of hedgehog in anterior imaginal disc structures in Drosophila results in upregulation of Ptc and increased growth and duplication of adult structures. Mutations in the human ptc gene are associated with hereditary basal cell nevus syndrome, which is associated with abnormal proliferation and patterning of many tissues.94 The mouse ptc gene is highly expressed in mesenchyme surrounding terminal lung buds. Over-expression of Shh from the SP-C enhancer/promoter in transgenic mice results in increased epithelial and mesenchymal proliferation, giving rise to a neonatal lethal lung phenotype which contains abundant mesenchyme and no functional alveoli.95 This is associated with upregulation of ptc expression, but not BMP-4, Wnt2 or FGF-7. Thus, Shh appears to play an important role as a mitogen for primitive respiratory epithelial cells in the growing tips of the early embryonic lung and regulates ptc expression in the adjacent mesenchyme.

Extracellular Matrix Components and Lung Morphogenesis During lung morphogenesis, proliferation and differentiation of epithelial and mesenchymal cells are accompanied by an active synthesis and deposition of extracellular matrix (ECM) proteins, including collagens, elastin, proteoglycans and glycoproteins.96,97 The ECM is principally deposited between epithelial and mesenchymal cells, forming the basement membrane, and around mesenchymal cells forming the interstitial matrix. Several studies have demonstrated that ECM components are differentially expressed, and have a specific cell distribution during lung morphogenesis. These observations suggest that ECM components may not only be a support of the tissue architecture, but may also play a direct role in the modulation of cell proliferation and cell differentiation. Absence or inhibition of the interaction of epithelial cells with the basement membrane has a direct consequence in the failure of normal lung development.10,11

Lung Epithelial Morphogenesis

227

Table 12.1. ECM components in the basement membrane during lung morphogenesis Component

Chain Composition

Mr (kDa)

Laminins LN 1 LN 2* LN 4* LN 6* LN 7*

heterotrimer (α1,β1,γ1) (α2,β1,γ1) (α2,β2,γ1) (α3,β1,γ1) (α3,β2,γ1)

400-840

Nidogen

monomer

150

Collagen IV

heterotrimer α1-α6(IV)

550

Proteoglycans (Perlecan)

monomer

SPARC

monomer

400-500 35

* Type of laminin variant distributed in the basement membrane during lung morphogenesis is proposed in accordance with the chain isoforms characterized. No purification of these LN variants from lung basement membrane has been reported (see text for details).

An extensive effort to characterize the different components localized in the basement membrane has taken place in the last decades.98-100 Table 12.1 shows a list of the principal basement membrane components found during embryonic lung development. Similarities exist with the composition of the basement membrane during lung morphogenesis that can also be found in other tissues. However, it is believed that differential expression of the ECM components and presence of specific isoforms are the key in the development of tissuefunction specificity. Laminins Laminins are a family of extracellular matrix glycoproteins which are composed of 3 chains, one central chain (α), and two lateral chains (β and γ) that are linked by disulfide bonds to form a cross-shaped molecule. To date, 5 α, 3 β and 2 γ chain isoforms have been characterized,101-105 resulting in the identification of ten variants of laminins (LN1 to LN10), that are distributed in a tissue specific manner.106 Only complete heterotrimers of LN are secreted and deposited in the basement membrane.107 The detection of specific chain isoforms by immunohistochemistry or molecular biology methods has been described during embryonic lung development.103,104,108-110 As shown in Table 12.1, the identification of these chain isoforms suggests the presence of five LN variants during lung development. So far, only the LN1 (α1,β1,γ1) variant has been extensively studied during lung branching morphogenesis in mammals.46,108,111-115 The importance and role of the other LN variants remains to be elucidated. LN is expressed in the earliest stage of embryonic development, indicating its important role in the organization of the basement membrane. Several reports have indicated that LN plays an important role in cell adhesion, migration, proliferation and differentiation during tissue development. During lung development, LN1 is expressed by epithelial and mesenchymal cells.108,111,112,114 Studies focused on the specific chain isoforms of LN have

228

Molecular Basis of Epithelial Appendage Morphogenesis

indicated that the α1 chain isoform is found principally deposited in the basement membrane at the epithelial-mesenchymal interface, with a predominant distribution in specific zones. Some studies have also indicated the presence of α1 around a few mesenchymal cells. The expression and distribution of the other two chains of isoform LN1, β1 and γ1, have a different pattern from α1. The localization of β1 and γ1 has been found in the basement membrane and around mesenchymal cells. Recent studies have determined that β1 and γ1 are constantly expressed during rabbit fetal lung development;108 meanwhile, the expression of the α1 chain has been found to increase at the end of fetal lung development.108,114 These findings show independent regulation of the expression of the chain isoforms. To elucidate the role of LN1 in lung morphogenesis, distinct domains of the three chain isoforms have been characterized.46,116,115 A domain in the cross region of the α1 chain is involved in the regulation of epithelial-mesenchymal interaction. In addition, the globular domains near the N-terminal of β1 and γ1 chains participate in the regulation of cell polarization. Recent studies in vitro have shown that LN1 has a role in promoting an organotypic rearrangement when isolated mesenchymal and epithelial cells from embryonic mouse lung are cultured together.46,114,115 The domain involved in this latter process has been localized to a globular region of the β1 chain. The identification of other chain isoforms, such as α2, has recently been described during lung development. The α2 chain isoform, found in LN2 (α2,β1,γ1) and LN4 (α2,β2,γ1) variants, is expressed during mouse and human lung development.101,110 In the preglandular stage of lung development, the α2 chain was observed to have a specific distribution around epithelial cells and smooth muscle cells in human110 and in mouse. In the canalicular stage in human fetal lung, α2 chain was present only in smooth muscle cells of bronchi.110 Studies in vitro, performed in our laboratory, have demonstrated that mesenchymal cells isolated from fetal mouse lung express the α2 chain isoform. In addition, LN2 plays a role in cell adhesion of these embryonic mesenchymal cells, being a better substrate for attachment than LN1. The α3 chain, found in LN6 (α3,β1,γ1) and LN7 (α3,β2,γ1) variants, has also been observed to be distributed in epithelial cells during fetal human lung development.110 The β2 chain isoform, found in the LN4 and LN7 variants, has been localized principally beneath epithelial cells.109 Further studies will be necessary to elucidate the biological role, and mechanism of action, of these chains during lung morphogenesis. Nidogen Nidogen is a 150 kDa glycoprotein synthesized principally by mesenchymal cells. Several studies have demonstrated that nidogen binds to the γ1 chain of LN, and forms a link between LN and Collagen IV.117,118 These findings suggest that nidogen has a key role in the organization of the basement membrane. Studies in lung morphogenesis have demonstrated that inhibition of the interaction of nidogen with LN blocks normal lung development.118-120 In addition, recent reports have shown that nidogen contains specific domains that are targets of proteinases, indicating that the susceptibility of nidogen to degradation may be a key step in the remodeling and degradation of the basement membrane.121,122 Collagen IV Collagen IV was one of the first ECM components to be identified in the basement membrane.123 It consists of three chains of 180 kDa bound in a triple helix structure. Collagen IV is an ubiquitous molecule, found in all basement membranes. It is frequently identified in a complex of four molecules, associated in a spider-shaped structure that facilitates the formation of a network structure. These observations suggest that collagen IV plays an important role in the structure of the basement membrane.

Lung Epithelial Morphogenesis

229

Recent studies of collagen IV chains have identified the presence of six chain isoforms in human, and five in mouse.124,125 Some reports have indicated that specific combinations of collagen IV chain isoforms follow a spatial and temporal distribution. It has been observed during kidney development that particular collagen IV combinations codistribute with specific LN variants. These observations strongly suggest that the specific expression of collagen IV molecules, together with LNs and nidogen, produce specific microenvironments in the basement membrane that may modulate cell migration, proliferation and differentiation. Collagen IV has been considered to be the main matrix scaffold in fetal lung basement membrane. Recent studies have identified the expression of different chain isoforms during lung development.126,127 However, the specific distribution and role of collagen IV molecules in the basement membrane during lung morphogenesis remains to be elucidated. Proteoglycans Proteoglycans comprise a core protein with sulfated carbohydrate chains.6,30 They function as flexible structures in the organization of the basement membrane and may also play an important role as a reservoir for growth factors, water and cations. Perlecan is a predominant proteoglycan in the basement membrane. It is composed of an approximate 450 kDa core protein with three heparan sulfate chains. Interaction of heparan sulfate proteoglycans with a specific domain of the β1 chain isoform of LN contribute directly in the organotypic arrangement.46 SPARC SPARC (secreted protein, acidic and rich in cysteine) is an anti-adhesive glycoprotein of the extracellular matrix. It has a wide distribution in the basement membrane during lung morphogenesis. Recent studies in which SPARC interactions were blocked by specific antibodies to this molecule indicate that it plays a role in lung branching morphogenesis.128

Physiological Determinants of Lung Morphogenesis Lung Liquid Distending Pressure Has Long Been Known to Play a Key Role in Determining Lung Growth, Yet the Molecular Mechanisms Connecting Fluid Pressure to Cellular Proliferation Are Largely Unknown The embryonic lung secretes fluid. The secretion of lung fluid is chloride dependent. Lung fluid production regulates lung growth, but does not appear to regulate branching per se.39 Increasing the distention pressure by increasing outflow resistance can partially reverse lung hypoplasia in experimental diaphragmatic hernia,129,130 and causes hyperplasia when the lung is initially normal.131 These data suggest that lung liquid volume regulation is crucial for “quantitative” lung growth,132 and that distension is linked to cell growth. Little is known about this except that IGF-II mRNA level varies inversely with lung volume.133,134 Recently, calcium entry through mechanical strain-activated ion channels has been shown to play a critical role in fetal lung epithelial cell proliferation.135 Activation of phospholipases and protein kinase C by PDGF-B and its receptor have also been implicated in the initiation of downstream strain-initiated events.136 EGF has also recently been shown to regulate both alveolar epithelial junctional permeability and active sodium transport as well as Na+, K+-ATPase α1 and β1 subunit expression.137,138 Interestingly, the α-amiloride-sensitive epithelial sodium channel -/- mouse dies in the neonatal period from failure to clear lung liquid,139 further confirming that factors that regulate lung liquid are essential both for normal lung development and for the smooth transition to air breathing. However, abrogation of chloride secretion into embryonic lung explants with chloride channel blocking diuretics decreased the size of the explants without decreasing the number of airway branches

230

Molecular Basis of Epithelial Appendage Morphogenesis

formed. Thus, chloride driven lung fluid may be important for lung growth, but not for branching morphogenesis per se. Clearly, over-production of lung liquid as can be stimulated by KGF, either in early embryonic mouse culture or in SP-C promoter-driven KGF over-expressing transgenic animals, results in deformation of the pulmonary epithelium similar to that found in the human lethal condition of cystic adenomatoid malformation.50,52,57

Hyperoxia, Even the “Natural Hyperoxia” Attending Human Premature Delivery, Has Long Been Considered to Be a Major Etiological Factor in the Adverse Impact of Premature Human Delivery on Lung Development Following acute hyperoxia in adult rodent models, the alveolar epithelium is denuded. Alveolar type 2 cells (AEC2) then regain the capacity to proliferate and repopulate the gas diffusion surface.140 Thus, in this special situation, AEC2 may be considered to have regained a stem cell function for alveolar epithelial repair.141 On the other hand, in neonatal rodents, acute hyperoxia results not only in proliferation of AEC2 to repair the denuded epithelium, but also in a significant inhibition of alveolar development.142 Alveolar hypoplasia is also an important sequela of neonatal hyperoxia both in primates and in human premature infants.143

Candidate Molecular Switches in the Transition from a Quiescent to a Proliferative Alveolar Epithelial Cell Phenotype and Back Again Following Acute Hyperoxia Include Autocrine Peptide Growth Factor Signaling Pathways and Cell Cycle Regulatory Elements TGF-β3 is the major TGF-β peptide secreted by rat AEC2 in culture, and it exerts an autocrine negative regulation on adult AEC2 proliferation.144 The production of TGF-β3 by these cells is dynamically downregulated during the proliferative phase of recovery from acute hyperoxia, but returns to normal levels following completion of recovery. Thus, the rate of DNA synthesis in rat AEC2 is inversely proportional to the autocrine production of TGF-β3.144 In the neonatal rat, hyperoxia results in a profound inhibition of alveolarization and is associated with high levels of TGF-β (mostly TGF-β1) activity in bronchial lavage. TGF-β1 levels and activity are also elevated in human premature neonatal lavage samples and presages the development of chronic lung disease. The majority of the latter TGF-β1 is probably derived from pulmonary macrophages.145 Therefore, we postulate that excess TGF-β1 production may play a key role in adversely regulating the normal temporo-spatial pattern of lung specific gene expression and morphogenesis (Fig. 12.5). We have also recently discovered that cyclin D2 and Cdc2 are specifically downregulated in quiescent adult AEC2.146 During the proliferative recovery phase following acute hyperoxia, these genes are reinduced, with activation of cdk4.141 These critical cell cycle control genes are also downregulated by TGF-β1 and cell-cell contact,146 further supporting a key role for TGF-β signaling in modulating the proliferative response to acute hyperoxia and the restoration of quiescence following recovery.

Perspectives on Lung Regeneration Novel Rational Pharmacological and Potential Gene Therapeutic Approaches to Modulating Lung Morphogenesis Organ regeneration has long been a biological reality in human skin wound healing and holds promise in human liver. In rodents, lung regeneration has also long been known to occur following lobectomy and is associated with coordinated cell proliferation, re-ex-

Lung Epithelial Morphogenesis

231

Fig. 12.5. A hypothesis diagram illustrating how physiological/environmental factors such as premature delivery, hyperoxia and ventilation could adversely impact the normal process of lung morphogenesis. Null mutation of the transcriptional factor Nkx-2.1, organ specific expression of a kinase deficient dominant negative form of FGFR, null mutation of EGFR, PDGFA, IGFR and of TGF-β3 all result in lethal abnormalities of lung morphogenesis. Therefore, it is inferred that normal function of these factors is necessary for normal lung morphogenesis. It is postulated that premature delivery, hyperoxia and mechanical ventilation induces expression of abnormally elevated levels of TGF-β1 and that excess TGF-β signaling adversely affects the function of the factors mentioned above. It is further hypothesized that abrogation of such abnormal TGF-β1 expression or signaling might ameliorate the adverse impact of these physiological/ environmental insults, mediated by excess TGF-β expression and signaling.

pression of elastin and formation of alveoli. However, whether lung regeneration can occur in primates or humans is controversial. Based on clinical experience with follow up of human premature infants with bronchopulmonary dysplasia, extensive lung growth and lung regeneration can occur naturally in these children. The issue of whether adult lung tissue, such as a parental lobe transplanted into a young person, can grow or regenerate remains to be settled, but the affirmative would be therapeutically desirable. Also, in cases of severe congenital pulmonary hypoplasia or dysplasia, for example diaphragmatic hernia and cystic adenomatoid malformation, lung regeneration could make the difference between nonsurvival and eventual recovery in many cases. In preliminary attempts to achieve the above desirable rational therapeutic goals, we have modeled the molecular structure of the TGF-β2 dimeric ligand interaction with the TGF-β I and II receptor ternary complex. This theoretical structure provides a rational framework for developing antibodies and cyclic peptides which can perturb TGF-β ligand-receptor interaction. Based on these structural biology computer modeling studies, we have devised an N-terminal TGF-β type II receptor peptide polyclonal antibody which perturbs specific features of TGF-β ligand-receptor interaction. This antibody abrogates TGF-β1 interaction with TGF-β type I receptors and perturbs interaction of ligand with the TGF-β type II receptor.147 The latter perturbation prevents assembly of ternary TGF-β type I and

232

Molecular Basis of Epithelial Appendage Morphogenesis

type II ligand-receptor signaling complexes. Preincubation of fetal mink lung epithelial cells with this antibody completely abrogates activation of the PAI-1 promoter by TGF-β1. But, inhibition of DNA synthesis by TGF-β1 is not adversely affected. Thus, the antibody appears to distinguish between TGF-β type I receptor mediated activation of extracellular matrix protein regulatory promoters and inhibition of the cell cycle through the TGF-β type II receptor. This antibody also has the interesting property of stimulating embryonic mouse lung morphogenesis in vitro.45 Our ultimate future aim is to devise new rational and gene therapeutic approaches to ameliorating lung injury and augmenting lung repair. The strategic approach we are following is therefore to develop rational therapeutic agents or gene therapy strategies that will regulate gene products or genes shown to play key roles in these processes. The ideal agent or agents would therefore mimic the instructive role of lung mesenchyme, and would correctly induce the temporo-spatial pattern of lung specific gene expression necessary to initiate and augment lung regeneration.

Acknowledgments The work in our laboratories is funded by the National Institutes of Health, HL 44060 and HL 44977 (DW) and by the Wellcome Trust (REO).

References 1. Comroe JH. Physiology of Respiration. Chicago: Year Book, 1965:11-16. 2. Morgan TH. Regeneration in Allolobophora feotida. Roux’s Arch Dev Biol 1897; 5:570-586. 3. Spemann H. Embryonic Development and Induction. New Haven: Yale University Press, 1938. 4. Zecca M, Basler K, Struhl G. Direct and long-range action of a wingless morphogen gradient. Cell 1996; 87:833-844. 5. Alescio T, Cassini A. Induction in vitro of tracheal buds by pulmonary mesenchyme grafted onto tracheal epithelium. J Exp Zool 1962; 150: 83-94. 6. Wessels NK. Mammalian lung development: Interactions in formation and morphogenesis of tracheal buds. J Exp Zool 1970; 175:455-466. 7. Shannon JM. Induction of alveolar type II cell differentiation in fetal tracheal epithelium by grafted distal lung mesenchyme. Devel Biol 1994; 166: 600-614. 8. Warburton D. Epigenetic autocrine and paracrine factors regulating lung morphogenesis. A paradigm for lung repair. Chest 1991;99:15S-18S. 9. Warburton D, Horcher P, Seth R et al. Epigenetic role of epidermal growth factor expression and signalling in embryonic mouse lung branching morphogenesis. Devel Biol 1992; 149:123-133. 10. Minoo P, King RJ. Epithelial-mesenchymal interactions in lung development. Ann Rev Physiol 1994; 56:13-45. 11. Hilfer SR. Morphogenesis of the lung: Control of embryonic and fetal branching. Ann Rev Physiol 1996; 58: 93-113. 12. Samakovlis C, Hacohen N, Manning G et al. Development of the Drosophila tracheal system occurs by a series of morphologically distinct but genetically coupled branching events. Development 1996; 122:1395-1407. 13. Wilk R, Weizman I, Shilo B-Z. Trachealess encodes a bHLH-PAS protein that is an inducer of tracheal cell fates in Drosophila. Genes & Development 1996; 10:93-102. 14. Isaac DD, Andrew DJ. Tubulogenesis in Drosophila: a requirement for the trachealess gene product. Genes & Development 1996; 10:103-117. 15. Andrew DJ, Horner MA, Pettit MG et al. Setting limits on homeotic gene function: Restraint of sex combs reduced activity by teashirt and other homeotic genes. EMBO J 1994; 13:1132-1144.

Lung Epithelial Morphogenesis

233

16. Sutherland D, Samakovis C, Krasnow MA. Branchless encodes a Drosophila FGF homologue that controls tracheal cell migration and the pattern of branching. Cell 1996; 67:1091-1101. 17. Peters K, Werner S, Liao X et al. Targeted expression of a dominant negative FGF receptor blocks branching morphogenesis and epithelial differentiation of mouse lung. EMBO J 1994; 13:3296-301. 18. Klambt C. The Drosophila gene pointed encodes two ETS-like proteins which are involved in the development of the midline glial cells. Development 1993; 117:163-176. 19. Guillemin K, Groppe J, Ducker K et al. The pruned gene encodes the Drosophila serum response factor and regulates cytoplasmic outgrowth during terminal branching of the tracheal system. Development 1996; 122:1353-1362. 20. Minoo P, Hamdan H, Bu D et al. TTF-1 regulates lung epithelial morphogenesis. Devel Biol 1995; 172:694-698. 21. Kimura S, Hara Y, Pineau T et al. The T/ebp null mouse: Thyroid-specific enhancer binding protein is essential for the organogenesis of the thyroid, lung, ventral forebrain and pituitary. Genes & Development 1996; 10:60-69. 22. Bohinski RJ, Huffman JA, Whitsett JA et al. Cis-active elements controlling lung cell-specific expression of human pulmonary surfactant protein B gene. J Biol Chem 1993; 268:11160-6. 23. Bohinski RJ, DiLauro R, Whitsett JA. The lung surfactant protein B gene promoter is a target for the thyroid transcription factor 1 and hepatocyte nuclear factor 3, indicating common factors for organ-specific gene expression along the foregut axis. Mol Cell Biol 1994; 14:5671-81. 24. Bruno MD, Bohinski RJ, Huelsman KM et al. Lung cell-specific expression of the murine surfactant protein A (SP-A) gene is mediated by interactions between the SP-A promoter and thyroid transcription factor-1. J Biol Chem 1995; 270:6531-6. 25. Ikeda K, Clark JC, Shaw-White JR et al. Gene structure and expression of human thyroid transcription factor-1 in respiratory epithelial cells. J Biol Chem 1995; 270:8108-8114. 26. Venkatesh VC, Planer BC, Schwartz M et al. Characterization of the promoter of human pulmonary surfactant protein B. Amer J Physiol Lung Cell Mol Biol 1995; 268:L674-82. 27. Yan C, Sever Z, Whitsett JA. Upstream enhancer activity in the surfactant protein B gene is mediated by thyroid transcription factor 1. J Biol Chem 1995; 270:24852-7. 28. Clevidence DE, Overdier DG, Peterson RS et al. Members of the HNF-3/forkhead family of transcription factors exhibit distinct cellular expression patterns in lung and regulate surfactant protein B promoter. Devel Biol 1994; 166:195-209. 29. Bingle CD, Gitlin JD. Identification of hepatocyte nuclear factor-3 binding sited in the Clara cell secretory protein gene. Biochem J 1993; 295:227-232. 30. Bingle CD, Hackett BP, Moxley M et al. Role of hepatic nuclear factor-3α and hepatocyte nuclear factor-3β in Clara cell secretory protein gene expression in the bronchiolar epithelium. Biochem J 1995; 308:197-202. 31. Jaskoll TF, Don-Wheeler G, Johnson R et al. Embryonic mouse lung morphogenesis and type II cytodifferentiation in serumless, chemically defined medium using prolonged in vitro cultures. Cell Diff 1988; 24:105-117. 32. Wuenschell CW, Sunday ME, Singh G et al. Embryonic mouse lung progenitor cells coexpress immunohistochemical markers of diverse mature cell lineages. J Histochem Cytochem 1996; 44:113-123. 33. Spooner BS, Hardman P, Paulsen A. Gravity in mammalian organ development: differentiation of cultured lung and pancreas rudiments during spaceflight. J Exp Zool 1994; 269:212-222. 34. Raaberg L, Nexo E, Buckley S et al. Epidermal growth factor transcription, translation and signal transduction in rat type II pneumocytes in culture. Amer J Resp Cell Mol Biol 1992; 6:44-49. 35. Warburton D, Lee M, Berberich MA et al. Molecular embryology and the study of lung development. Minisymposium Report, Cell & Developmental Biology Branch, Lung Division, NHLBI, NIH and The Society for Developmental Biology. Amer J Resp Cell Mol Biol 1993; 9:5-9.

234

Molecular Basis of Epithelial Appendage Morphogenesis

36. Kaartinen V, Voncken JW, Shuler CA et al. Abnormal lung development and cleft palate: defects of mesenchymal-epithelial interaction in mice lacking TGF-β3. Nature Genetics 1995; 11:415-421. 37. Miettinen P, Berger JE, Meneses J et al. Epithelial immaturity and multiorgan failure in mice lacking epidermal growth factor receptor. Nature 1995; 376:337-41. 38. Raaberg L, Nexo E, Jorgensen PE et al. Fetal effects of epidermal growth factor deficiency in rats by autoantibodies against epidermal growth factor. Pediatr Res 1995; 37:175-181. 39. Souza P, Kuliszewski M, Wang J et al. PDGF-AA and its receptor influence early lung branching via an epithelial-mesenchymal interaction. Development 1995; 121:2559-67. 40. Shiratori M, Oshika E, Ling PU et al. Keratinocyte growth factor and embryonic lung morphogenesis. Amer J Resp Cell Mol Biol 1996; 15:328-338. 41. Seth R, Shum L, Wu F et al. Role of epidermal growth factor expression in early mouse embryo lung branching morphogenesis in culture: Antisense oligodeoxynucleotide strategy. Devel Biol 1993; 158: 555-559. 42. Serra R, Pelton RW, Moses HL. TGF-β1 inhibits branching morphogenesis and N-Myc expression in lung bud organ cultures. Development 1994; 120:2153-2161. 43. Serra R, Moses HL. pRb is necessary for inhibition of N-Myc expression by TF-β1 in embryonic lung organ cultures. Development 1995; 121:3057-3066. 44. Zhao Y, Young SL. Expression of transforming growth factor-beta type II receptor in rat lung is regulated during development. Amer J Physiol Lung Cell Mol Physiol 1995; 269:L419-426. 45. Zhao JS, Bu D, Lee MK et al. Abrogation of TGF- type II receptor stimulates embryonic mouse lung morphogenesis. Devel Biol 1996; 180:242-257. 46. Schuger, L, Skubitz APN, Gilbride K et al. Laminin and heparan sulfate proteoglycan mediate epithelial cell polarization in organotypic cultures of embryonic lung cells: Evidence implicating involvement of the inner globular region of laminin B1 chain and the heparan sulfate groups of heparan sulfate proteoglycan. Devel Biol 1996; 179:264-273. 47. Raaberg L, Nexo E, Poulsen SS et al. An immunologic approach to induction of epidermal growth factor deficiency: induction and characterization of autoantibodies to epidermal growth factor in rats. Pediatr Res 1995a; 37:169-174. 48. Raaberg L, Nexo E, Jorgensen PE et al. Fetal effects of epidermal growth factor deficiency induced in rats by autoantibodies against epidermal growth factor. Pediatr Res 1995b;37:175-181. 49. Korfhagen TR, Swantz RJ, Wert SE et al. Respiratory epithelial cell expression of human transforming growth factor-α induces lung fibrosis in transgenic mice. J Clin Invest 1994; 93:1691-1699. 50. Simonet WS, Derose ML, Bucay N et al. Pulmonary malformation in transgenic mice expressing human keratinocyte growth factor in the lung. Proc Nat Acad Sci USA 1995; 92:12461-12465. 51. Shiratori M, Oshika E, Ung LP, Singh G, Shinozuka H, Warburton D, Michalopoulos G, Katyal SL. Keratinocyte growth factor and embryonic rat lung morphogenesis. Am J Respir Cell Mol Biol 1996; 15:328-338. 52. Zhou L, Graeff RW, McCray PB et al. Keratinocyte growth factor stimulates CFTR-induced fluid secretion in the fetal lung in vitro. Amer J Physiol Lung Cell Mol Physiol 1996; 15:L987-994. 53. Deterding RR, Shannon RM. Proliferation and differentiation of fetal rat pulmonary epithelium in the absence of mesenchyme. J Clin Invest 1995; 95:2963-2972. 54. Nogawa H, Ito T. Branching morphogenesis of embryonic mouse lung epithelium in mesenchyme-free culture. Development 1995; 121:1015-1022. 55. Panos RJ, Bak PM, Simonet WS et al. Intratracheal instillation of keratinocyte growth factor decreases hyperoxia-induced mortality in rats. J Clin Invest 1995; 96:2026-2033. 56. Sonnenberg E, Weidner KM, Birchmeier C. Scatter factor/hepatocyte growth factor and its receptor, the c-Met tyrosine kinase, can mediate a signal exchange between mesenchyme and epithelia during mouse development. J Cell Biol 1993; 123:223-235.

Lung Epithelial Morphogenesis

235

57. Shiratori M, Michalopoulos G, Shinozuka H et al. Hepatocyte growth factor stimulates DNA synthesis in alveolar epithelial type II cells in vitro. J Respir Cell Mol Biol 1995; 12:171-180. 58. Hepatocyte growth factor/scatter factor induces a variety of tissue-specific morphogenetic programs in epithelial cells. J Cell Biol 1995; 13:1573-1586. 59. Batchelor DC, Hutchins AM, Klempt M et al. Developmental changes in the expression patterns of IGFs, type 1 IGF receptor and IGF-binding proteins-2 and-4 in perinatal rat lung. J Mol Endocrinol 1995; 15:105-115. 60. Lallemand AV, Ruocco SM, Joly PM et al. In vivo localization of the insulin-like growth factors I and II (IGF I and IGF II) gene expression during human lung development. Int J Devel Biol 1995; 39:529-537. 61. Maitre B, Clement A, Williams MC et al. Expression of insulin-like growth factor receptors 1 and 2 in the developing lung and their relation to epithelial cell differentiation. Amer J Resp Cell Mol Biol 1995; 13:262-270. 62. Rechts-Bogart GZ, Moats-Stats BM, Howard K et al. Cellular localization of messenger RNAs for insulin-like growth factors (IGFs), their receptors and binding proteins during fetal rat lung development. Amer J Resp Cell Mol Biol 1996; 14:61-69. 63. Schuller AG, van Neck JW, Beukenholdt RW et al. IGF, type I IGF receptor and IGF-binding protein mRNA expression in the developing mouse lung. J Mol Endocrinol 1995; 14:349-355. 64. Liu JP, Baker J, Perkins SA et al. Mice carrying null mutations of the genes encoding insulin-like growth factor I (Igf-1) and type 1 IGF receptor (IGF-IR). Cell 1993; 75:59-72. 65. Coppola D, Ferber A, Miura M et al. A functional insulin-like growth factor I receptor is required for the mitogenic and transforming activities of the epidermal growth factor receptor. Mol Cell Biol 1994; 14:4588-4595. 66. Buch S, Jassal D, Cannigia I et al. Ontogeny and regulation of platelet-derived growth factor gene expression in distal fetal rat lung epithelial cells. Amer J Resp Cell Mol Biol 1994; 1:251-261. 67. Bostrom H, Willetts K, Pekny M et al. PDGF-A signaling is a critical event in lung alveolar myofibroblast development and alveogenesis. Cell 1996; 85:863-873. 68. Aguayo SM, Schuyler WE, Murtagh JJ et al. Regulation of lung branching morphogenesis by bombesin-like peptides and neutral endopeptidase. Amer J Resp Cell Mol Biol 1994; 10:635-642. 69. Pelton RW, Dickinson ME, Moses HL et al. In situ hybridization analysis of TGF-β 3 RNA expression during mouse development: Comparative studies with TGF-β 1 and 2. Development 1990; 110:609-620. 70. Moens CB, Stanton BR, Parada LF et al. Defects in heart and lung development in compound heterozygotes for two different targeted mutations at the N-Myc locus. Development 1993; 119:485-499. 71. Wang J, Kuliszewski M, Yee W et al. Cloning and expression of glucocorticoid-induced genes in fetal rat lung fibroblasts. Transforming growth factor-β3. J Biol Chem 1995; 270:2722-8. 72. Bellusci S, Henderson R, Winnier G et al. Evidence from normal expression and targeted misexpression that bone morphogenetic protein (BMP-4) plays a role in mouse embryonic lung morphogenesis. Development 1996; 122:1693-1702. 73. Heine UI, Munoz EF, Flanders KC. Colocalization of TGF-β1 and collagen I and II, fibronectin and glycosaminoglycans during lung branching morphogenesis. Development 1990; 109:29-36. 74. Murthy US, Anzano MA, Stadel JM et al. Coupling of TGF-beta-induced mitogenesis to G-protein activation in AKR-2B cells. Biochem Biophys Res Commun 1988; 152:1228-1235. 75. Kataoka R, Sherlock J, Lanier SM. Signaling events initiated by transforming growth factor-beta 1 that require Gi alpha 1.J Biol Chem 1993; 268:19851-19857. 76. Wang T, Donahoe PK, Zervos AS. Specific interaction of type I receptors of the TGF-β family with the immunophilin FKBP-12. Science 1994; 265:674-676. 77. Chen RH, Miettinen PJ, Maruoka EM et al. A WD-domain protein that is associated with and phosphorylated by the type II TGF-β receptor. Nature 1995; 377:548-552.

236

Molecular Basis of Epithelial Appendage Morphogenesis

78. Yamaguchi K, Shirakabe K, Shibuya H et al. Identification of a member of the MAPKKK family as a potential mediator of TGF-β mediated signal transduction. Science 1995; 270:2008-2011. 79. Sekelsky JJ, Newfeld SJ, Raftery LA et al. Genetic characterization and cloning of mothers against dpp, a gene required for decapentaplegic function in Drosophila melanogaster. Genetics 1995; 139:1347-1358. 80. Liu F, Hata A, Baker JC et al. A human Mad protein acting as a BMP-regulated transcriptional activator. Nature 1996; 381:620-623. 81. Newfeld SJ, Chartoff EH, Graff JM et al. Mothers against dpp encodes a conserved cytoplasmic protein required in DPP/TGF-beta responsive cells. Development 1996; 122:2099-2108. 82. Wiersdorff V, Lecuit T, Cohen SM et al. Mad acts downstream of Dpp receptors, revealing a differential requirement for dpp signaling in initiation and propagation of morphogenesis in the Drosophila eye. Development 1996; 122:2153-2162. 83. Zhang Y, Feng X, We R et al. Receptor-associated Mad homologues synergize as effectors of the TGF-β response. Nature 1996; 383:168-172. 84. Hildebrand A, Romaris M, Rasmussen LM et al. Interaction of the small interstitial proteoglycans biglycan, decorin and fibromodulin with transforming growth factor-β. Biochem J 1994; 302:527-534. 85. Yamashita H, Ichijo-H, Grimsby S et al. Endoglin forms a heteromeric complex with the signaling receptors for transforming growth factor-β. J Biol Chem 1994; 269:1995-2001. 86. Lastres P, Letamendia A, Zhang H et al. Endoglin modulates cellular responses to TGF-β1. J Cell Biol 1996; 133:1109-1121. 87. McAllister KA, Grogg KM, Johnson DW et al. Endoglin, a TGF-β binding protein of endothelilal cells is the gene for hereditary hemorrhagic telangiectasia type 1. Nature Genetics 1994; 8:345-351. 88. Heberlien U, Wolff T, Rubin GM. The TGF homolog Dpp and the segment polarity gene hedgehog are required for propagation of a morphogenetic wave in the Drosophila retina. Cell 1993; 75:913-926. 89. Basler K, Struhl G. Compartment boundaries and the control of Drosophila limb pattern by hedgehog protein. Nature 1994; 368: 208-214. 90. Capdevilla J, Pariente F, Sampedro J et al. Subcellular localization of the segment polarity gene patched suggests an interaction with the wingless reception complex in Drosophila embryos. Development 1994; 120:987-998. 91. Diaz-Benjumea FJ, Cohen B, Cohen SM. Cell interaction between compartments establishes the proximal-distal axis of Drosophila legs. Nature 1994; 372: 175-179. 92. Bitgood MJ, McMahon AP. Hedgehog and BMP genes are co-expressed at many diverse sites of cell-cell interaction in the mouse embryo. Dev Biol 1995; 172: 126-138. 93. Urase K, Mukasa T, Irigashi H et al. Spatial expression of Sonic hedgehog in the lung epithelium during branching morphogenesis. Biochem Biophys Res Commun 1996; 225:161-166. 94. Johnson RL, Rothman AL, Xie J et al. Human homolog of patched, a candidate gene for the basal cell nevus syndrome. Science 1991; 272:1668-1671. 95. Bellusci S, Yasuhide F, Rush MG et al. Involvement of Sonic hedgehog (Shh) in mouse embryonic lung growth and morphogenesis. Development 1997; 124:53-63. 96. Dunsmore SE, Rannels DE. Extracellular matrix biology in the lung. Amer J Physiol Lung Cell Mol Physiol 1996; 270: L3-L27. 97. McGowan ES. Extracellular matrix and the regulation of lung development and repair. FASEB J 1992; 6: 2895-2904. 98. Paulsson M. Basement membrane proteins: Structure, assembly and cellular interactions. Crit Rev Biochem Mol Biol 1992; 27, 93-127. 99. Timpl R. Macromolecular organization of basement membrances. Curr opin Cell Biol 1996; 8:618-624. 100. Timpl R, Brown JC. Supramolecular assembly of basement membranes. Bioessays 1996; 18, 123-132.

Lung Epithelial Morphogenesis

237

101. Bernier SM, Utani A, Sugiyama S et al. Cloning and expression of laminin α 2 chain (Mchain) in the mouse. Matrix Biol 1994; 14: 447-455. 102. Galliano M, Aberdam D, Aguzzi A et al. Cloning and complete primary structure of the mouse α3 chain. J Biol Chem 1995; 270:21821-21826. 103. Iivanainen A, Sainio K, Sariola H et al. Primary structure and expression of a novel human laminin α4 chain. FEBS Letters 1995; 365:183-188. 104. Iivanainen A, Vuolteenaho R, Sainio et al. The human laminin β2 chain (S-laminin): Structure, expression in fetal lung tissues and chromosomal assignment of the LAMB2 gene. Matrix Biol 1995; 14: 489-497. 105. Vuolteenaho R, Nissinen M, Sainio K et al. Human laminin M chain (Merosin): Complete primary structure, chromosomal assignment, and expression of the M and A chain in human fetal tissues. J Cell Biol 1994; 381: 381-394. 106. Burgerson RE, Chiquet M, Deutzmann R et al. A new nomenclature for the laminins. Matrix Biol 1994; 14: 209-211. 107. De Arcangelis A, Neuville P, Boukamel R et al. Inhibition of laminin α1-chain expression leads to alteration of basement membrane assembly and cell differentiation. J Cell Biol 1996; 133: 417-430. 108. Durham PL, Snyder JM. Characterization of alpha 1, beta 1, and gamma 1 laminin subunits during rabbit fetal lung development. Dev Dyn 1995; 203:408-421. 109. Durham PL, Snyder JM. Regulation of the beta 2 subunit chain of laminin in developing rabbit fetal lung tissue. Differentiation 1996; 60:229-243. 110. Virtanen I, Laitinen A, Tani T et al. Differential expression of laminins and their integrin receptors in developing and adult human lung. Amer J Resp Cell Mol Biol 1996; 15:184-196. 111. Klein G, Ekblom M, Fecker L et al. Differential expression of laminin A and B chains during development of embryonic mouse organs. Development 1990; 110: 823-837. 112. Lallemand AV, Ruocco SM, Gaillard DA. Synthesis and expression of laminin during human foetal lung development. Anat Rec 1995; 242:233-241. 113. Schuger L, O’Shea KS, Nelson BB et al. Organotypic arrangement of mouse embryonic lung cells on a basement membrane extract: involvement of laminin. Development 1990; 110:1091-1099. 114. Schuger L, Varani J, Killen PD et al. Laminin expression in the mouse lung increases with development and stimulates spontaneous organotypic rearrangement of mixed lung cells. Devel Dynam 1992; 195:43-54. 115. Schuger L, Skubitz APN, Morenas A et al. Two separate domains of laminin promote lung organogenesis by different mechanisms of action. Dev Biol 1995; 169:520-532. 116. Matter ML, Laurie GW. A novel laminin E8 cell adhesion site required for lung alveolar formation in vitro. J Cell Biol 1994; 124:1083-1090. 117. Reinhardt D, Mann K, Nischt R et al. Mapping of nidogen binding sites for collagen type IV, heparan sulfate proteoglycan, and zinc. J Biol Chem 1993; 268: 10881-10887. 118. Dziadek M. Role of laminin-nidogen complexes in basement membrane formation during embryonic development. Experientia 1995; 51:901-913. 119. Ekblom P, Ekblom M, Fecker L et al. Role of mesenchymal nidogen for epithelial morphogenesis in vitro. Development 120, 2003-2014, 1994. 120. Senior RM, Griffin GL, Mudd MS et al. Entactin expression by rat lung and rat alveolar epithelial cells. Amer J Resp Cell Mol Biol 1996; 14:239-247. 121. Mayer U, Mann K, Timpl R et al. Sites of nidogen cleavage by proteases involved in tissue homeostasis and remodeling. Eur J Biochem 1993; 217:877-884. 122. Alexander CM, Howard EW, Bissell MJ et al. Rescue of mammary epithelial cell apoptosis and entactin degradation by a tissue inhibitor of metalloproteinases-1 transgene.J Cell Biol 1996; 135:1669-1677. 123. Kefalides NA. Structure and biosynthesis of basement membrane. Int Rev Connect Tiss Res 1973; 6:63-71. 124. Hudson BG, Reeders ST, Tryggvason K. Type IV collagen: structure gene organization, and role in human diseases. J Biol Chem 1993; 268:26033-26036.

238

Molecular Basis of Epithelial Appendage Morphogenesis

125. Miner JH, Sanes JR. Collagen IV α3, α4, and α5 chains in rodent basal laminae: Sequence, distribution, association with laminins, and developmental switches. J Cell Biol 1994; 127:879-891. 126. Chen J, Little CD. Cellular events associated with lung branching morphogenesis including the deposition of collagen type IV. Dev Biol 1987; 120:311-321. 127. Thomas T, Dziadek M. Expression of collagen 1(IV), laminin and nidogen genes in the embryonic mouse lung: Implications for branching morphogenesis. Mech Dev 1994; 45:193-201. 128. Strandjord TP, Sage EH, Clark JG. SPARC participates in the branching morphogenesis of developing fetal rat lung. Amer J Resp Cell Mol Biol 1995; 13:279-287. 129. Hedrick HL, Kaban JM, Pacheco BA et al. Prenatal glucocorticoids improve pulmonary morphometrics in fetal sheep with congenital diaphragmatic hernia. J Pediatr Surg 1997; 32:217-221. 130. Wilson JM, DiFiore JW, Peters CA. Experimental fetal tracheal ligation prevents pulmonary hypoplasia associated with fetal nephrectomy: Possible application of congenital diaphragmatic hernia. J Pediatr Surg 1993; 28:1433-1440. 131. Moessinger AC, Harding R, Adamson TN et al. Role of lung fluid volume in growth and maturation of the fetal sheep lung. J Clin Invest 1990; 86:1270-1277. 132. Hopper SB, Harding R. Fetal lung liquid: A major determinant of the growth and functional development of the fetal lung. Clin Exp Pharmacol Physiol 1995; 22:235-247. 133. Harding R, Hooper SB, Han VKM. Abolition of fetal breathing movements by spinal cord transection leads to reductions in fetal lung liquid volume, lung growth and IGF-II gene expression. Pediatr Res 1993; 34:148-153. 134. Hooper SB, Wallace MJ, Harding R. Amiloride blocks the inhibition of fetal lung liquid secretion caused by AVP but not by asphyxia. J Appl Physiol 1993; 74:111-115. 135. Liu M, Xu J, Tanswell AK, Post M. Inhibition of mechanical strain-induced fetal rat lung cell proliferation by gadolinium, a stretch-induced channel blocker. J Cell Physiol 1994; 161:50-507. 136. Liu M, Liu J, Buch S, Tanswell AK et al. Antisense oligonucleotides for PDGF-B and its receptor inhibit mechanical strain-induced fetal lung cell growth. Amer J Physiol Lung Cell Mol Physiol 1995; 269:L178-84. 137. Borok Z, Hami A, Danto SI et al. Effects of EGF on alveolar epithelial junctional permeability and active sodium transport. Amer J Physiol Lung Cell Mol Physiol 1996; 270:L559-565. 138. Danto SI, Zabski SM, Borok Z et al. Regulation of Na+, K+ -ATPase α-1 and β-1 subunit protein expression by epidermal growth factor. Amer J Resp Crit Care Med 1995; 151:A190. 139. Hummler E, Barker P, Gatzy J et al. Early death due to defective neonatal lung liquid clearance in αENAC-deficient mice. Nature Genetics 1996; 12:325-328. 140. Crapo JD, Barry BE, Foscue HA et al. Structural and biochemical changes in rat lungs occurring during exposures to lethal and adaptive levels of oxygen. Amer Rev Resp Dis 1980; 122:123-143. 141. Bui KC, Buckley S, Wu F et al. Induction of A and D type cyclins and cdc2 kinase activity during recovery from short term hyperoxic lung injury. Amer J Physiol Lung Cell Mol Biol 1995; 268:L625-635. 142. Blanco LN, Frank L. The formation of alveoli in rat lung during the third and fourth postnatal weeks: Effects of hyperoxia, dexamethasone and deferrioxamine. Pediatr Res 1993; 34:334-340. 143. deLemos RA, Coalson JJ. The contribution of experimental models to our understanding of the pathogenesis and treatment of bronchopulmonary dysplasia. Clin Perinatol 1992; 19:521-539. 144. Buckley S, Bui KC, Hussain M et al. Dynamics of TGF-β3 peptide activity during alveolar epithelial type 2 cell proliferative recovery from acute hyperoxic injury in the rat. Amer J Physiol Lung Cell Mol Physiol 1996; 271:L54-60.

Lung Epithelial Morphogenesis

239

145. Kotecha S, Wangoo A, Silverman M et al. Increase in the concentration of transforming growth factor β-1 in bronchoalveolar lavage fluid before development of chronic lung disease of prematurity. J Pediatr 1996; 128:464-469. 146. Wu F, Buckley S, Bui KC et al. Cell-cycle arrest in G0/G1 phase by contact inhibition and transforming growth factor-β1 in mink Mv1Lu epithelial cells. Amer J Physiol Lung Cell Mol Physiol 1996; 270:L879-888. 147. Hall FL, Benya PD, Padilla S et al. TGF-β type II receptor signalling: Investigations of intrinsic/associated casein kinase activity, receptor interaction, and functional effects of blocking antibodies. Biochem J 1996; 316:303-310.

Part IV

Molecular Mechanism

CHAPTER 13

Early Molecular Events in Feather Morphogenesis: Induction and Dermal Condensation Randall B. Widelitz and Cheng-Ming Chuong

Introduction

E

mbryogenesis is composed of a series of inductive events that change the fate of responding tissues. Through induction, new tissues and organs are generated. In the beginning of embryogenesis, ectodermal-endodermal interactions produce the mesoderm by what is termed primary induction. In the past ten years, rapid progress in the molecular cascades involved in primary induction has been made, particularly in Xenopus.1,2 Later in development, many tissues and organs arise from epithelial-mesenchymal interactions through secondary inductions. As shown by many chapters in this book, molecular cascades involved in these epithelial-mesenchymal interactions are just starting to be revealed. It is interesting that, so far, several key molecules responsible for primary inductive events also function during secondary inductions (see chapters 6 for hair, 9 for teeth; 14 for teeth; and 12 for lung). We have been studying the molecular basis for secondary inductions using feather morphogenesis as the model. The chicken feather model is well suited to this purpose because: 1. it is accessible to experimental manipulation including recombination and transplantation; 2. it can be reliably cultured as an in vitro explant; 3. many feather buds develop over the surface of the body with a growth gradient; 4. rich amounts of data generated from classical experiments provide an insightful understanding to the process of epithelial-mesenchymal interactions;3 and 5. nonlethal mutations are available. Feathers grow from the interactions of flat epithelium and mesenchyme into an organized array of epithelial appendages (Fig. 13.1). Neither the epithelium nor the mesenchyme can form the skin appendages by itself. Without reciprocal interactions, new feathers will not form and existing feathers will disintegrate.4,5 Early in development the mesenchyme acquires inductive properties and determines the number, size, location and structural identities of the appendage, while the responding epithelium determines the orientation and competence state, and modulates responses under developmental constraints (summarized in refs. 3, 6-9). However, the molecular cascade responsible for these processes remains to be worked out. In chapter 5 an overview of the different phases of feather morphogenesis was discussed. The goal of this chapter is to focus on the current status of molecules known Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

244

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 13.1. Formation of individual feather primordia from a homogeneous feather field. Interactions between the epithelium and mesenchyme of an initially homogeneous piece of skin (A) from a stage 29 embryo induce dermal condensations which develop into feather buds (B) after 2 days in culture.

to be involved in two of the early phases: induction and dermal condensation. The formation of the primordia involves initiation, propagation and termination of propagation. The precise location of primordia initiation sites is a question related to pattern formation. Models theorizing how this may be established are discussed in chapter 18. In this chapter we will focus on molecules experimentally essential for initiation or suppression of initiation. We will then review molecules involved in mesenchymal condensation, and how the size of mesenchymal condensation, or the future diameter of the appendage, may be regulated.

Molecules in the Feather Inductive Phase Feather induction arises at stage 28 along the midline primary row of the lumbo-sacral region and progresses in a posterior to anterior direction. Adjacent rows subsequently develop bilaterally.6,10-12 The feathers become arranged in an organized and well characterized hexagonal pattern. This pattern has facilitated the identification of molecules involved in feather morphogenesis, including growth factors, signaling molecules, transcription factors and adhesion molecules. Recently, new progress has been made on early molecular events that lead to the induction of skin appendage development. By identifying molecular signals that regulate the induction process, we can begin to examine events preceding the early morphological changes associated with feather formation and other structures derived from epithelial-mesenchymal interactions.

Early Molecular Events in Feather Morphogenesis

245

What are the initial inductive signals required for feather induction? One straightforward approach to identifying the responsible molecular pathway is to determine the expression sequence of candidate molecules. However, in actual practice, this has been difficult to assess, since embryos selected at the same times may vary in developmental age, and feather morphogenesis progresses faster than the resolution of embryonic staging.13 We have done our best to order several candidate molecules using this approach (please see chapter 20). Another approach to examining the expression sequence is to use a feather regeneration assay in which the epithelium and mesenchyme are separated and then recombined. Under these experimental conditions, feather development is reset and the reappearance of important regulatory molecules can be determined. Using this approach, we have determined that FGF-4 and bone morphogenetic proteins (BMPs) -2 and -4 do not disappear following epithelial-mesenchymal separation. Sonic hedgehog (Shh) and Wnt7a disappear in 3 h and reappear at 6 h. Notch, Serrate, and Delta reappear around 10 h. Msx-1 and Msx-2 reappear at 18 h. Then neural cell adhesion molecules (NCAM) and homeobox proteins such as HoxC6 become localized to the anterior bud.5,14 The order of reappearance of gene expression in regenerating feather buds may or may not parallel their sequence during initial development, but gives a perspective of a functional molecular cascade. In the future we will use double in situ hybridization to compare the sequence of molecular expression during normal development in higher resolution. Meanwhile, we will now discuss the expression patterns of these molecules and their demonstrated function in feather development. We will also attempt to explore their interrelationships during normal feather development to establish a tentative molecular model for the formation of feather buds. The heterogeneous distribution of many molecules at the asymmetric short feather bud stage (see chapter 5 for stage definition) is schematically represented in Figure 13.2.

Fibroblast Growth Factors (FGFs) FGFs are a family of peptide growth factors that stimulate cell proliferation, morphogenesis and wound repair. At least ten different FGF isoforms have been identified.15 FGFs play a role in the primary induction of mesoderm16 and in secondary inductions such as in limb buds,17-20 teeth (chapters 9, 14) and lungs (chapter 12). Several lines of evidence suggest that FGFs act as endogenous inducers of feather formation. They are localized within the skin before feather formation. FGF-4 is initially expressed as a continuous epithelial stripe along the dorsal midline at stage 28, later segregating into the placodes of the developing feathers.21 In lateral rows, FGFs such as FGF-4 first appear in the epithelial placodes.22 The FGF-2 expression pattern has not been characterized prior to stage 33, but it is expressed in the placode epithelium at this time.23 FGF-8, on the other hand, is not expressed in the skin during early feather induction (our unpublished data). To further assess the roles of some of these molecules we have applied them ectopically and examined the ensuing morphological and molecular changes. When FGF-1 (500 ng/ml) and FGF-2 (500 ng/ml) were added to the culture media of stage 31 skin explants, adjacent feather buds fused. The interbud regions were apparently converted into bud regions. At stage 34, exogenous FGF-1 and -2 increased the number but decreased the size of new lateral edge feather buds. Moving toward the midline, FGF treatment also produced a dense cell region devoid of feathers near the lateral edge. Still closer to the midline, FGFs induced feather buds that were wider and shorter than control buds.24 An example showing the effects of FGF-2 is shown (Fig. 13.3B). The differential response may be due to the competence state of the responding skin. To test the effects of local FGF applications, Affigel blue beads were soaked in FGFs -1, -2 or -4 (850 ng/ml) for one hour at 37°C (using the method described in Hayamizu et al25) and applied as a local source to the dorsal skin. The FGFs

246 Fig. 13.2. Molecular heterogeneity of feather buds. Schematic diagram profiling the heterogeneous distribution of key molecules in the late stage of short feather buds. Before the formation of feather buds, these molecules are either homogeneously distributed or absent. The different modes by which they become restricted to the feather bud domain or interbud domain are shown in Figure 5. (See insert for color.)

Molecular Basis of Epithelial Appendage Morphogenesis

FGFR1 FGFR2 FGF-4

BMP-2 BMP-4

Shh Msx

TGFβ-2 Type II TGF βR

PKC PKA

Early Molecular Events in Feather Morphogenesis

247

induced feathers in competent epithelium, even within regions of the skin that normally remain apteric.24 Scaleless is an avian autosomal recessive mutant with mostly naked skin. No scales form, and a few abnormal feathers are found near the midline and femoral regions.26 Recombination studies showed that the original defect is in the epithelium, and that the mesenchyme is originally normal. Failing to receive an appropriate message from the epithelium, the mesenchyme subsequently becomes abnormal.27 It was recently shown that FGFs can rescue this molecular defect by inducing new feather buds from the skin of the scaleless mutant.23 This suggests that in the molecular cascade of induction, FGF acts downstream to the putative scaleless product. To examine the relationship with other molecules of interest, we treated skin prepared from stage 28 embryos with FGF-4 (850 ng/ml) and then looked at the distribution of other molecules by in situ hybridization. Our preliminary data showed that, at 16 h, FGF-4 inhibited the expression of FGF-4 transcription, possibly through feedback inhibition, while causing BMP-4 and Shh expression to become more diffuse. The difficulty here is to pinpoint the time of maximal alteration, and we may have to complete a time course study to see the whole picture of molecular modulation. If FGFs can stimulate feather initiation, then FGF receptors (FGFRs) must be present in the skin. Four FGFRs have been identified with different binding affinities for each of the ten FGF isoforms. They have been localized to the developing skin.23,28 FGFR1 is found in the dermal condensations. FGFR2 is in the anterior epithelial placode and in the interbud mesenchyme. FGFR3 is found throughout the bud and interbud mesenchyme. FGFR4 has not yet been mapped within the developing chicken skin. More work needs to be done to complete our understanding of how FGF-receptor interactions regulate feather induction. However, the FGF receptors are known to act as tyrosine kinases.29 The phosphorylation cascade acts through Grb in association with Sos-1 to activate Ras, then Raf, Mek and finally Erk.30-32 Grb, Ras, Raf and Erk-1 are enriched in the dermis beneath the feather buds but not in the interbud regions,24 supporting the notion that the FGF pathway is involved in the induction of feather buds.

Sonic Hedgehog (Shh) Shh is a secreted growth factor that is a member of the hedgehog growth factor family.33 It is secreted in two forms exerting both short and long range effects on tissue patterning.34 Shh is expressed in vertebrate tissues with inductive and polarizing activities.33,35-37 It plays a role in mesenchymal cell proliferation in the lung38 and in neural floor plate induction.39 In the chicken limb, Shh regulates the expression of the Gli and Gli3 transcription factors.40 In chicken dorsal skin, Shh first is expressed as a stripe along the midline at stage 28. The continuous linear pattern segregates into individual feather buds at stage 29. Expression then propagates bilaterally by stage 33.21 Shh expression in the lateral rows initiates in the epithelial placodes. It is expressed distally in the midline epithelial placodes of stage 35 embryos.22,41 Epithelial-mesenchymal recombination studies showed that mesenchymal factors are required to maintain Shh expression in the placode. Ectopic expression of Shh from the RCAS replication competent retroviral vector42 to the epithelium and mesenchyme induced dermal condensations with a large accumulation of mesenchymal cells.41 These condensations may be induced by stimulated cell migration and/or cell proliferation. Recently, K14-Shh was shown to induce multiple foci of basal cell carcinoma and activated mesenchyme in transgenic mice.43 Shh probably plays a role in active epithelial-mesenchymal interactions. When the events are regulated properly, a new skin appendage forms. If not, overexpression of Shh leads to deregulated new growth, such as tumors.

248

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 13.3. Effects of exogenous growth factors on feather bud formation. Growth factors were added to the culture media of developing stage 29-31 dorsal embryo skin. (a) Control, (b) FGF-2, (c) BMP-2.

a

Ctrl b

FGF-2 c

BMP-2

Early Molecular Events in Feather Morphogenesis

249

We also examined the effects of ectopic Shh expression on other molecules of interest. Ectopic Shh expression induced TGF-β2 and inhibited PKC activity,41 consistent with the notion that Shh favors the formation of feather primordia.

Patched Patched is a target of Shh41,44 and may function as its receptor in conjunction with smoothened.45,46 In Drosophila patched suppresses transcription. When hedgehog binds to patched it relieves the suppression.47 Mutations of the patched gene have been associated with the formation of basal cell carcinomas,48 which is mimicked by the overexpression of Shh in transgenic mice.43 In chicken skin, patched is also expressed along the midline at stage 28 and becomes discontinuous by stage 29 with expression in the feather bud but not the interbud domain.21 Patched is initially expressed by the epithelium at stage 28, but its expression shifts to the mesenchyme by stage 31.

Bone Morphogenetic Proteins (BMPs) BMPs are members of the TGF superfamily. They were initially identified for their effects on osteocyte proliferation and differentiation (reviewed in ref. 49). BMPs were subsequently found to affect other developmental systems in a myriad of ways. BMPs are ventralizing agents in Xenopus embryos whose activity is blocked by chordin and noggin.50 However, heterodimers of BMP-4 and -7 induce mesoderm in Xenopus embryos.51 Genetic knockout experiments also showed that BMP-4 is required for mouse mesodermal induction.52 BMPs are involved in tooth morphogenesis.53,54 In the limb, Shh and FGF-4 induce the expression of BMP-2 which is involved in limb patterning.55,56 In the gut Shh can also induce the expression of BMP-4.57 BMP-4 induces the expression of transcription factors Msx-1, Msx-2 and Egr-1 in the tooth (chapter 14). BMPs are related to the Decapentaplegic (Dpp) gene in Drosophila. Dpp also is involved in tissue induction and patterning.58 Therefore, BMPs and related proteins are involved in tissue induction and patterning in many different systems. We examined the distribution of the BMPs in developing skin. BMPs -2 and -4 are not present in the early skin. BMP-2 appears around the time of epithelial placode formation, initially appearing toward the center of the mesenchymal condensation.21 Later expression moves toward the anterior mesenchyme and expands to include the anterior epithelium. BMP-4 initially appears in the feather primordia at the time of epithelial placode formation. It is expressed in the epithelium and mesenchyme with the highest expression at the epithelial-mesenchymal border. Later, expression moves toward the anterior mesenchyme.21 The role of BMPs was explored further by adding exogenous BMPS to skin explants.21 When added to the media of cultured explants, BMPs-2 and -4 (100-1000 ng/ml) did not affect the more mature, midline feather buds, but increased the spacing of the newer, lateral edge feathers with dose-dependence, suggesting that they act early in the feather formation pathway. An example of the effects of BMP-2 is shown (Fig. 13.3c). Local effects were determined by placing beads soaked in BMP-2 or BMP-4 (33 µg/ml–1 mg/ml) on explant cultures. Surrounding each bead was a zone of feather inhibition. The size of the zone was concentration-dependent.21 While FGFs act as positive regulators to induce feather formation, we were surprised to find that BMPs act as negative regulators to inhibit feather formation, since their expression is initiated within the feather bud and not in the interbud space. This information may contribute to the development of models for periodic pattern formation (chapter 18). The effects of BMPs on other molecules were determined by in situ hybridization. After a 16 h exposure, both BMP-2 (500 µg/ml) and BMP-4 (337 µg/ml) inhibited the expression

250

Molecular Basis of Epithelial Appendage Morphogenesis

of BMP-2, BMP-4, Fgf-4 and Shh transcripts.21 This feedback inhibition may contribute to feather patterning through lateral inhibition. BMPs bind to heterodimers of type I and type II serine/threonine kinase BMP receptors (BMPRs). Binding to the receptor transmits a signal through transcription factors that are homologs of the mother’s against Decapentaplegic (Mad) gene product found in Drosophila. In chicken embryos, BRK-1 and -2 are type I receptors, while BRK-3 and -4 are type II receptors.59 Not much has been done to determine the distribution of the BMPRs in chicken skin development. Expression of a dominant negative type I BMPR in hind limb buds transformed some scales into feather-type scales.60 To study the specific roles of the BMP pathway in different types of skin appendages, it would be interesting to compare the distribution of BMPs and their receptors in feathers and scales.

Wnt Genes Wnt genes are homologs of the Drosophila wingless gene and are secreted peptide growth factors believed to function through the frizzled receptor. Wnt7a is important for axis determination in the chicken limb. The limbs of transgenic mice with targeted disruption of the Wnt7a gene become ventralized,61 suggesting that Wnts are involved in establishing the dorsal-ventral limb pattern. Ectopic expression of Wnts in the ventral hemisphere of Xenopus embryos induces axis duplication.62,63 Other Wnt gene expression antagonizes glycogen synthase kinase 3β, which normally phosphorylates adenomatous polyposis coli (APC) to target β-catenin for degradation. Therefore, Wnt can expression can increase cellular β-catenin pools, which interact with LEF-1/TcF and regulate nuclear gene expression.64 In the skin, Wnt-11 is found localized within the mesenchyme of feathers at the epithelial-mesenchymal border of stage 35-36 chicken embryos.65 We have found that at a similar stage, Wnt7a is expressed in the posterior placode epithelium. Using a skin recombination assay, we found that Wnt7a expression in the epithelium was mesenchyme-dependent.5 Given the importance of Wnt pathways in other development systems, we expect that future work will show the key roles they play in feather morphogenesis.

Msx Genes Msx-1 and -2 are homologs of the Drosophila msh genes often found in the context of epithelial—mesenchymal interactions, suggesting that they are involved in inductive processes. The Msx genes encode homeobox transcription factors.66 In Xenopus, Msx-1 functions as a ventralizing agent and is induced by BMP-4.67 BMPs also have been shown to induce Msx genes in other developmental systems.68-70 Msx genes are involved in the proper formation of many developmental structures. Transgenic mice with targeted disruption of Msx-1 have craniofacial abnormalities effecting the skeleton, tooth eruption and hair formation.71 Transgenic mice overexpressing Msx-2 from a CMV promoter showed hyperkeratotic skin lesions and abnormal hair phenotypes.72,73 In other developmental systems, Msxs and FGFs have been shown to regulate the expression of one another,74 and are likely to be involved in the same molecular pathway. Expression of a mutated type II FGF receptor mimics the craniofacial abnormalities found with disrupted Msx-1.75 In stage 31 embryonic skin, Msx-1 is expressed in the feather placode epithelium. Its expression becomes asymmetric, favoring the anterior epithelium at stage 33. Expression extends proximally, alleviating the asymmetry by stage 35.76 Msx-2, on the other hand, is expressed in the distal epithelial placode of stage 31 chicken embryo skin with some weak mesenchymal staining. Expression spreads anteriorly by stage 33. At stage 35 expression is in the distal half of the epithelium with some mesenchymal staining remaining.76 Both Msx genes are only expressed in the feather bud domains, not in the interbud domains. The

Early Molecular Events in Feather Morphogenesis

251

function of the Msx genes in feather induction is not entirely understood, but they may play a role in regulating proliferation/differentiation.76

Tissue-Nonspecific Alkaline Phosphatase (TNAP) TNAP is anchored to cell membranes by a phosphatidylinositol linkage.77 In chicken skin, it is first expressed as a stripe along the dorsal midline at stage 29. Later this changes to a periodic pattern as the expression becomes restricted to the feather buds. Inhibiting TNAP during early feather bud formation results in delayed feather bud formation and causes dose-dependent feather fusion.78 Later inhibition did not affect the initiated midline feather buds, but inhibited the latent feather buds in the lateral portions of the skin. While a specific role for TNAP in feather patterning has not been established, its early expression preceding feather formation and the developmental delays caused by its inhibition strongly suggest that it is involved in establishing the distribution of feathers within the feather field. It is not known whether TNAP is acting as an enzyme or structural protein in this process.78

Molecules in the Dermal Condensation Phase Mesenchymal condensation, the accumulation of mesenchymal cells around sites of activated epithelia, is a fundamental process at all sites of epithelial-mesenchymal interactions. It is the site of active morphogenesis and remodeling. Many signaling molecules, adhesion molecules, proteases and inhibitors, etc. are expressed in regions of mesenchymal condensation. In the skin, cell density rises from 2.6 nuclei/1000 µm3 in the homogeneous dense dermis to 5.2 nuclei/1000 µm3 within the dermal condensations.3 Is this due to cell redistribution, proliferation, or both? Proliferation has been reported to stop for about 24 h during dermal condensation,79 so cell migration probably plays a major role, particularly in the early phase of dermal condensation formation. This is coupled by the finding that hexagonally arranged collagen fibrils are found in the dense dermis in early morphogenetic stages. These fibrils appear to work as “cellular highways” for cell migration leading to the feather primordia localized at the fibril junctions.80 The scaleless mutant chicken, mentioned above, fails to form these dermal collagen fibrils.81 Disruption of collagen synthesis with β-aminopropionitrile also inhibits feather formation.82

Extracellular Matrix Molecules Proteoglycan, collagen, tenascin, fibronectin Blocking proteoglycan synthesis with para-nitrophenyl-β-D-xyloside (2mM) disrupts normal feather patterning, showing the importance of adhesion to extracellular matrix molecules in the formation of feather dermal condensations.83 Extracellular matrix molecules such as collagen and fibronectin are expressed in exquisite patterns and may provide tracks on which cells migrate.80,84 Tenascin is an extracellular matrix protein85 containing three domains resembling epidermal growth factor, fibronectin type II and fibrinogen.86 It functions as a hexamer.87 Tenascin has been shown to promote cell migration and mesenchymal condensation during development.68,88,89 Tenascin is enriched in developing feathers and may be involved in their formation.90 During skin morphogenesis, tenascin is restricted to the anterior placode epithelium by the feather placode stage and spreads slightly to the mesenchyme by early feather bud formation. Inhibition of tenascin activity with an anti-tenascin antibody caused the feather buds to remain round and stop growth. These phenotypes are similar to that produced by cAMP91 and they may belong to the same molecular pathway. Fibronectin is an extracellular matrix protein which binds to several integrin receptors, primarily via an Arg-Gly-Asp peptide sequence. Its expression has been associated with cell

252

Molecular Basis of Epithelial Appendage Morphogenesis

migration, proliferation, differentiation and apoptosis.92 During early skin morphogenesis, fibronectin is widespread throughout the mesenchyme.90 To test the function of fibronectin in feather morphogenesis, skin explants from stage 33 embryos were incubated with antifibronectin antibodies. Inhibition of fibronectin activity did not affect feathers growing along the midline that had entered a more mature stage. However, treatment caused the epithelium to separate from the mesenchyme toward the lateral edge of the explants, preventing further development of these young buds.90

Cell Adhesion Molecules NCAM, DCC, integrin Cell-cell adhesion is required for proper dermal condensation formation. The neural cell adhesion molecules (NCAM) are members of the immunoglobulin family, mediating cell-cell adhesion. NCAM first appears at around stage 29 at the basal level in the dense dermis. NCAM then becomes enriched in the dermal condensations of stage 35 embryonic skin93 and disappears from the interbud regions. Inhibition of NCAM with anti-NCAM antibodies perturbed the formation of dermal condensations. Some condensations were large, some were small and some were missing.90 The effect occurred with dose-dependence. Each condensation which formed could elongate normally with proper orientation. The expression pattern and antibody inhibition results suggest that NCAM is involved in dermal condensation formation of skin appendages. DCC was first identified as a gene deleted in colorectal carcinoma.94 It is a member of the immunoglobulin family of adhesion molecules with close amino acid homology to NCAM and neogenin.95,96 Recently, it was found to bind to netrin-1,97-99 which guides the migration of neurons.100-103 Netrin contains laminin homology and can promote or inhibit growth cone migration. In the skin, DCC is present in embryonic epithelium, including epidermis and endoderm.104 Antibody inhibition of DCC activity in developing skin from stage 33 chicken embryos blocked the formation of dermal condensations and left the dermal cells diffusely distributed. This suggests that proper function of the DCC pathway is required for epithelial-mesenchymal interactions underlying the formation of dermal condensations. In this capacity, DCC may function between epithelial cells or between epithelial cells and their substrate. Integrins are transmembrane adhesion molecules which function as dimers of α and β subunits.105 The composition of their α and β subunits determines their binding affinity for fibronectin, collagen, laminin, vitronectin, etc. To date, about 20 different α and β combinations have been identified. Integrins are involved in cell migration, proliferation, differentiation and signal transduction. They also help to organize actin microfilament structure.106 In the feather, integrin β1 is expressed in the epithelial cells and localizes to the epithelialmesenchymal interface of the developing placode, but is not present in the interbud space.90 Inhibiting integrin activity in skin from stage 33 embryos with an anti-integrin β1 antibody resulted in the separation of the epithelium from the mesenchyme in younger feather buds and inhibited their further development. Treatment of older feather buds with anti-integrin β1 blocked their elongation and they remained short and round.

Extracellular Signaling Molecules TGF-β and Shh The TGF-β family of growth factors regulates growth control in many experimental systems.107 Depending on the target cells, they stimulate or inhibit cell proliferation. They

Early Molecular Events in Feather Morphogenesis

253

also regulate cell migration. In addition, TGF-β regulates the expression of other downstream molecules such as adhesion molecules and extracellular matrix molecules.108,109 We explored the role of TGF-β in skin appendage formation. In situ hybridization shows that TGF-β2 is expressed uniformly in the bud and interbud epithelium of placode stage feathers.110 By stage 35, TGF-β2 mRNA becomes restricted to the feather epithelium and mesenchyme. TGF-β2 protein is enriched at the epithelial-mesenchymal interface of the distal feather buds.111 The role of TGF-β2 in dermal condensations was tested by placing Affigel blue beads soaked in TGF-β2 (7.5 µg/ml) on cultured skin explants. Under these conditions, TGF-β2 treatment induced a ring of tightly spaced feather buds.111 TGF-β1 at similar doses did not affect these cultures, nor did TGF-β2 placed in the culture medium at the concentration of 3-30 ng/ml. A larger effect was produced by sandwiching the TGF-β2 coated bead between the epithelium and mesenchyme, following epithelial-mesenchymal separation and recombination. In fact, TGF-β2 coated beads can stimulate dermal condensation formation from denuded mesenchyme in the absence of epithelium. Within a limited range, the size of the condensations increases with the dose of TGF-β2. To test whether these condensations were capable of inducing feather buds, the growth factor beads were removed after 24-36 h and replaced with epithelium from stage 31-33 embryos. Many feather buds formed on top of the condensations formed by TGF-β2 treatment, but only rarely outside the TGF-β2 treated regions.111 Consistent with the previous finding, addition of TGF-β2 induced the expression of tenascin-C and NCAM, molecules shown to be involved in the formation of dermal condensation. There are three types of TGF-β receptors.112 TGF-β binds to the type II and type III receptors. The type III receptor lacks the kinase domain and is anchored to the membrane via a proteoglycan. It probably regulates the availability of TGF-β for binding to the type II receptors. The type II receptor binds to TGF-β and subsequently forms a complex with the type l receptor. After complex formation, the type I receptor signals through a kinase domain. Here we examined the distribution of TGF-β type II receptor and found that it is distributed throughout the bud and interbud epithelium and mesenchyme at stage 31. It then becomes localized to the bud domain around stage 34-35. When TGF-β2 coated beads were placed on top of naked mesenchyme, the TGF-β receptor was induced in the absence of epithelium, perhaps through a positive feedback loop.111 Our preliminary data showed that antibody to type II β receptor increases the spacing between feather buds. Besides its role in feather induction, Shh is expressed in the epithelium prior to dermal condensation formation.41,113 Transduction of the RCAS retrovirus directing the constitutive (LTR promoter) expression of Shh produced enlarged dermal condensations which seemed to form by fusing neighboring feather fields to form a gigantic feather buds.41 Examination of these buds showed a wider distribution of TGF-β2, NCAM, and suppression of PKC (see below). These events may contribute to the enhanced formation of dermal condensations. Phosphorylation of CREB (which also favors the formation of dermal condensation, see below) does not change.

Intracellular Signaling Molecules PKA and PKC The above sections have dealt with the role of growth factors and their receptors in feather morphogenesis. Growth factors bind to their receptors at the cell surface, yet these interactions produce changes elsewhere within the cell. The action of growth factors is translated at the cell surface and transmitted by a cascade of molecular signals. We have begun to explore how these signals affect feather formation. Among these molecules, the expression

254 Fig. 13.4. The size of feather bud domain and the ratio of activators and inhibitors. We propose that the size and placement of feather primordia are modulated by the relative amount of activators (light gray) and inhibitors (dark gray) (left column), viewed as longitudinal sections. The Y axis shows the expression levels. The right column shows the corresponding size of the feather bud domain (black) and interbud spacing (white) viewed from the top. When the effective concentration of the activator(s) is much greater than that of the inhibitor(s), then a big feather bud will form. In regions where the effective concentration of the inhibitor(s) is higher than that of the activator(s), an interbud space forms. Thus, the effective amount of ligands and receptors in different skin regions can determine the size and number of feather buds. (See color insert).

Molecular Basis of Epithelial Appendage Morphogenesis

Side View

Top View

Positive signal

Feather domain

Negative signal

Interbud domain

patterns of the phosphorylated cAMP responsive element binding protein (pCREB) and protein kinase C (PKC) are strikingly complementary. Initially PKC (using a pan-PKC antibody that recognizes all classical forms of PKC, UBI—Lake Placid, NY) appears in the mesenchyme immediately subjacent to the epithelium at stage 30. PKC spreads throughout the dermis by stage 31. As the feather begins to form at stage 32, PKC starts to disappear from the feather mesenchyme but remains in the interbud regions. PKC continues to disappear from the feather mesenchyme but stays in the interbud regions by stage 34. CREB, the unphosphorylated cAMP responsive element binding protein, is expressed ubiquitously in the epithelium and mesenchyme at all stages of feather development. When CREB is activated, it is converted to pCREB by activated kinases. We used antibodies that can differentiate the two forms to examine the distribution of CREB and pCREB. At stage

Early Molecular Events in Feather Morphogenesis

255

29, there is no pCREB. Gradually, as placodes form, pCREB appears in the epithelium and mesenchyme within the bud region, but not the interbud region.91 To further examine the role of these signaling molecules, PKA and PKC levels were modulated with specific agonists and antagonists. PKC activity was stimulated in explants prepared from stage 34 embryos by exposure to 4 β-phorbol-12,13-diacetate (100 nM). This treatment produced small, widely dispersed buds.91 Higher concentrations (1 µM) produced total inhibition of feather formation. Inhibition of PKC activity with Calphostin C (20-250 nM) increased the size of feathers slightly. Activation of PKC also leads to the suppression of NCAM. In contrast, treatment of stage 34 skin with forskolin (20 µM), an activator of adenylyl cyclase, or 8(4CPT)-cAMP, a cAMP analog, increased the size and fusion of dermal condensations in stage 30-31 skin. In later skin, they caused feather buds to remain symmetric and blocked elongation. It seems that increasing cAMP concentrations caused the dermal cells to condense so tightly that the buds failed to progress beyond this developmental stage. This hypothesis is supported by the inhibition of Shh, Notch, Delta and Serrate expression within these cultures. Shh may be required for elongation of the long bud.114 Notch, Delta and Serrate appear to be involved in the formation of anterior-posterior asymmetry formation.14 The activators for PKA and PKC and how these signals link to feather induction remain to be determined. Cellular oncogenes In an early stage of dermal condensation formation, cell proliferation stops for approximately 24 h in the center of dermal condensations.79 Cell proliferation then increases in the late stage of dermal condensation formation to drive the growth of feathers from the short bud to the long bud stage. This can be seen by the increased BrdU labeling in the bud region. Increased expression of some cellular oncogenes coincides with this increase of cell proliferation. c-Myc is a transcription factor which forms a complex with Max. It acts to regulate cell proliferation and apoptosis. In the feather, c-Myc appears in the epithelial placode but not in the interbud epithelium.115 c-Ets, a nuclear transcription factor which regulates metalloprotease expression is present in the mesenchyme of developing feather buds.115 Inhibition of c-Ets with hydrocortisone inhibited type IV collagenase activity and inhibited feather formation.116

Size Determination: Boundary and Spacing Several molecules that favor dermal condensation are discussed in the above section. The question arises: What molecules regulate the size of dermal condensations? Secreted factors, such as TGF-β2 and Shh would have an effect over the broadest area and could act as morphogens to regulate dermal condensation size. Activin, a secreted factor, has been shown to exert an effect at a distance of 10 cells.117 The distance of influence may differ for other secreted factors. Transcription factors, signaling molecules and adhesion molecules would act more locally. Since Shh was shown to induce the expression of TGF-β2, they may function through a common pathway. The gradient of TGF-β2 expression may be converted to a sharp boundary between the area of dermal condensation and the interbud region if a threshold level is required for TGF-β2 activity (reviewed in ref. 118). The difference of feather size is apparent in different feather tracts. Do different sized feathers start with the same primordium size and then grow into different sized feathers? Or, do they initially start with different primordium sizes? Observations of spinal, wing and caudal feathers suggest that both factors may contribute to feather size. From their initial appearance, the diameter of the flight feather primordia is larger than the small feather primordia in the spinal tract. Subsequent, daily elongation rates are also different (see below).

256

Molecular Basis of Epithelial Appendage Morphogenesis

What are the critical factors that determine the diameter of a feather primordium? Is it the concentration of cells (number of cells), the concentration of morphogens, or the total mass of cells that matters? In the mature follicles, it has been shown that the size of the hairs is proportional to the size of the dermal papilla. Again, whether it is the number of dermal papilla, or the concentration of growth factors, that matters has not been determined. What is the molecular nature of the “intrinsic property” that determines the size of the primordia? We propose that the ratio of activated activator and inhibitor pathways are likely to play an important role (Fig. 13.4). The strength of the activator or inhibitor pathway is determined by the concentration of ligands and the amount of receptors and signaling molecules downstream to receptors. From the experimental data so far, we suggest that FGF/Shh are the activators, and BMP-2/BMP-4 are the inhibitors. The combination of activators and inhibitors not only determines the site of initiation, but their relative strengths also regulate the boundary of the dermal condensations.

Expression Modes and Molecular Cascades If the spatial distribution of signaling molecules and adhesion molecules can determine the location and size of feather bud domains, then it is important to know how these molecules reach that particular distribution pattern. Therefore we analyze the temporal and spatial changes of these molecules and hope to deduce some testable hypothesis for molecular interactions. Analysis of the activator and inhibitor expression patterns in stage 28 to 32 embryos revealed at least four different categories. These four modes of molecular localization to their respective sites in the skin are presented (Fig. 13.5). Two modes apply to the activators and two modes apply to the inhibitors. Mode 1. Some activators initiate with a homogeneous distribution at low expression levels which segregate into discrete units as the molecules become enhanced in the bud region and suppressed in the interbud region. Examples are NCAM and TGF-β2. Mode 2. A second scenario starts with a complete absence of expression; the activator first appears in a periodic pattern in the primordial region. FGF, Shh and pCREB are in this category. Mode 3. Some inhibitors start their expression homogeneously throughout the dense dermis. As the primordia start to form, their expression begins to disappear from the primordial region, but remains in the interprimordia region. An antibody to pan-PKC showed that PKC proteins fall within this category. Mode 4. Other inhibitors are not initially expressed, but first appear in a periodic pattern, in the formed primordia but not in the interprimordia region. This distribution is counter-intuitive for an inhibitor and suggests that a reaction-diffusion mechanism is likely to be involved. BMP-2 and BMP-4 belong to this category. Further analyses of the sequence of these expression modes may help us to identify and arrange molecular and cellular events in induction and dermal condensations.

Summary What begins to emerge is a chronology of molecular cascades mediating epithelialmesenchymal interactions to regulate feather induction and dermal condensation formation. With the above modes, the earliest expressed molecules are the homogeneously distributed NCAM, TGF-β, PKC etc. Later, as they become restricted to their specific locations, precisely determining whether their expression pattern becomes restricted earlier or later than the direct appearance of FGF and Shh in the feather primordia is difficult. BMPs are expressed slightly later. The balance achieved by these molecules will sharpen the TGF-β expressing domain. TGF-β and its receptor also have a self-augmenting mechanism to sharpen the border of dermal condensations. Within the dermal condensation, expression of tenascin

Early Molecular Events in Feather Morphogenesis

Fig. 13.5. Schematic diagram showing different modes of molecular expression during dermal condensation. The top bar represents regions of feather bud domain (striped rectangle) and interbud domain (open rectangle), viewed as longitudinal sections. The Y axis shows expression levels in a scale from 0-100%. Steady state expression levels of skin prior to feather formation are shown as the dotted line. All the molecules are originally homogeneously distributed, either at high levels (at the level of 100), medium levels (at 50), or absent (at 0). They become expressed specifically in the bud domain or interbud domain through different modes. To study the mechanism, we have identified four different expression modes: (1) Some activators, such as TGF-β2 and NCAM become expressed by cells within the feather. Those closest to the center of the feather exhibit the highest expression levels, and the levels of expression fall off, with the minimum expression level falling toward the middle of the interbud space. (2) Other activators, such as FGF, Shh and pCREB are not expressed prior to feather formation. Their expression is initiated in a defined region within the feather bud. (3) Some inhibitors such as PKC are initially expressed uniformly through the bud and interbud region but their expression levels decrease toward the center of the bud region. (4) The expression of other inhibitors, such as BMPs, is initiated toward the center of the forming bud. We presume that they only inhibit in the interbud region, where their effective concentration is higher than that of the activator. The different modes are likely the results of different molecular interactions during early feather morphogenesis. A tentative working model of these interactions is shown in Fig. 13.6.

257

258

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 13.6. A proposed working model for the molecular and tissue interactions underlying feather morphogenesis. From the effect of the molecules on feather bud formation, the order of molecular expression, and effect of one molecule on the expression of other molecules, we suggest a tentative molecular cascade.

and NCAM is increased. Both adhesion molecules are required for mesenchymal cell migration and condensation. Studies on the relationship of these molecules showed that ectopic Shh expression induces the formation of dermal condensations and the expression of TGF-β2.41 Placement of TGF-β2 beads on skin mesenchyme explants induces the expression of tenascin in the absence of epithelium.111 Similarly, NCAM mediates dermal condensation formation and is induced by TGF-β2.111 We also find that TGF-β2 beads can inhibit the expression of PKC, an inhibitor of feather formation.111 More relationships between these important molecules are being studied. A tentative sequence and the epithelial (E) or mesenchymal (M) origin of the molecules in this pathway is shown in Figure 13.6. We hope it will serve as a framework for the placement of new molecules and processes involved in this process.

Acknowledgment The authors wish to thank Dr. Jiang for contribution to Figure 13.3. This work was supported by grants from NIH (CMC, RW), Norris Cancer Center (RW), NSF (CMC), and CTR (CMC).

References 1. Smith JC. Mesoderm-inducing factors and mesodermal patterning. Curr Opin Cell Biol 1995; 7:856-861. 2. Kessler DS, Melton DA. Vertebrate embryonic induction: Mesodermal and neural patterning. Science 1994; 266:596-604. 3. Sengel P. Morphogenesis of skin. Cambridge: Cambridge University Press, 1976. 4. Rawles ME. Tissue interactions in scale and feather development as studied in dermalepidermal recombination. J Embryol Exp Morph 1963; 11:765-789. 5. Chuong C-M, Ting-Berreth S, Widelitz RB et al. Early events during the regeneration of skin appendages: Order of molecular reappearance following epithelial-mesenchymal recombination with rotation. J Invest Dermatol 1996; 107:639-646.

Early Molecular Events in Feather Morphogenesis

259

6. Linsenmayer TF. Control of integumentary patterns in the chick. Dev Biol 1972; 27:244-271. 7. Novel G. Feather pattern stability and reorganization in cultured skin. J Embryol exp Morph 1973; 30:605-633. 8. Reynolds AJ, Jahoda CA. Cultured dermal papilla cells induce follicle formation and hair growth by transdifferentiation of an adult epidermis. Development 1992; 115:587-593. 9. Chuong CM. The making of a feather: Homeoptroteins, retinoids and adhesion molecules. Bioessays 1993; 15:513-521. 10. Mayerson PL, Fallon JF. The spatial pattern and temporal sequence in which feather germs arise in the white leghorn chick embryo. Dev Biol 1985; 109:259-267. 11. Davidson D. The mechanism of feather pattern development in the chick. 1. The time of determination of feather position. J Embryol exp Morph 1983a; 74:245-259. 12. Davidson D. The mechanism of feather pattern development in the chick. II. Control of the sequence of pattern formation. J Embryol Exp Morph 1983b; 74:261-273. 13. Hamburger V, Hamilton HL. A series of normal stages in development of the chick embryo. J Morphol 1951; 88:49-91. 14. Chen C-WJ, Jung H-S, Jiang T-X et al. Asymmetric expression of Notch/Delta/Serrate is associated with the anterior-posterior axis of feather buds. Dev Biol 1997; 188:181-187. 15. Ohuchi H, Nakagawa T, Yamamoto A et al. The mesenchymal factor, FGF10, initiates and maintains the outgrowth of the chick limb bud through interaction with FGF8, an apical ectodermal factor. Development 1997; 124:2235-2244. 16. Slack JM, Isaacs HV, Song J et al. The role of fibroblast growth factors in early Xenopus development. Biochem Soc Symp 1996; 62:1-12. 17. Niswander L, Tickle C, Vogel A et al. FGF-4 replaces the apical ectodermal ridge and directs outgrowth and patterning of the limb. Cell 1993; 75:579-587. 18. Fallon JF, Lopez A, Ros MA et al. FGF-2: Apical ectodermal ridge growth signal for chick limb development. Science 1994; 264:104-107. 19. Cohn MJ, Izpisua-Belmonte JC, Abud H et al. Fibroblast growth factors induce additional limb development from the flank of chick embryos. Cell 1995; 80:739-746. 20. Crossley PH, Minowada G, MacArthur CA et al. Roles for FGF8 in the induction, initiation, and maintenance of chick limb development. Cell 1996; 84:127-136. 21. Jung H-S, Francis-West PH, Widelitz RB et al. Local inhibitory action of Bmps and interfeather bud spacing: A model for periodic pattern formation. Dev Biol 1998; 196:11-23. 22. Nohno T, Kawakami Y, Ohuchi H et al. Involvement of the Sonic hedgehog gene in chick feather formation. Biochem Biophys Res Commun 1995; 206:33-39. 23. Song H, Wang Y, Goetinck PF. Fibroblast growth factor 2 can replace ectodermal signaling for feather development. Proc Natl Acad Sci USA 1996; 93:10246-10249. 24. Widelitz RB, Jiang T-X, Noveen A et al. FGF induces new feather buds from developing avian skin. J Invest Dermatol 1996; 107:797-803. 25. Hayamizu TF, Sessions SK, Wanek N et al. Effects of localized application of transforming growth factor beta 1 on developing chick limbs. Dev Biol 1991; 145:164-173. 26. Abbott UK, Asmundson VS. Scaleless, an inherited ectodermal defect in the domestic fowl. J Hered 1957; 48:63-70. 27. McAleese SR, Sawyer RH. Correcting the phenotype of the epidermis from chick embryos homozygous for the gene scaleless (sc/sc).Science 1981; 214:1033-1034. 28. Noji S, Koyama E, Myokai F et al. Differential expression of three chick FGF receptor genes, FGFR1, FGFR2 and FGFR3, in limb and feather development. Prog Clin Biol Res 1993; 383B:645-654. 29. Fernig DG, Gallagher JT. Fibroblast growth factors and their receptors: An information network controlling tissue growth, morphogenesis and repair. Prog Growth Factor Res 1994; 5:353-377. 30. Roberts TM. Cell biology. A signal chain of events. Nature 1992; 360:534-535. 31. McCormick F. How receptors turn Ras on. Nature 1993; 363:15-16. 32. Marshall CJ. Specificity of receptor tyrosine kinase signaling: Transient versus sustained extracellular signal-regulated kinase activation. Cell 1995; 80:179-185.

260

Molecular Basis of Epithelial Appendage Morphogenesis

33. Riddle RD, Johnson RL, Laufer E et al. Sonic hedgehog mediates the polarizing activity of the ZPA. Cell 1993; 75:1401-1416. 34. Bumcrot DA, Takada R, McMahon AP. Proteolytic processing yields two secreted forms of sonic hedgehog. Mol Cell Biol 1995; 15:2294-2303. 35. Echelard Y, Epstein DJ, St-Jacques B et al. Sonic hedgehog, a member of a family of putative signaling molecules, is implicated in the regulation of CNS polarity. Cell 1993; 75:1417-1430. 36. Krauss S, Concordet JP, Ingham PW. A functionally conserved homolog of the Drosophila segment polarity gene hh is expressed in tissues with polarizing activity in zebrafish embryos. Cell 1993; 75:1431-1444. 37. Levin M, Johnson RL, Stern CD et al. A molecular pathway determining left-right asymmetry in chick embryogenesis. Cell 1995; 82:803-814. 38. Bellusci S, Furuta Y, Rush MG et al. Involvement of Sonic hedgehog (Shh) in mouse embryonic lung growth and morphogenesis. Development 1997; 124:53-63. 39. Ericson J, Morton S, Kawakami A et al. Two critical periods of Sonic Hedgehog signaling required for the specification of motor neuron identity. Cell 1996; 87:661-673. 40. Marigo V, Johnson RL, Vortkamp A et al. Sonic hedgehog differentially regulates expression of GLI and GLI3 during limb development. Dev Biol 1996a; 180:273-283. 41. Ting-Berreth SA, Chuong C-M. Sonic hedgehog in feather morphogenesis: Induction of mesenchymal condensation and association with cell death. Dev Dyn 1996a; 207:157-170. 42. Hughes SH, Greenhouse JJ, Petropoulos CJ et al. Adaptor plasmids simplify the insertion of foreign DNA into helper-independent retroviral vectors. J Virol 1987; 61:3004-3012. 43. Oro AE, Higgins KM, Hu Z et al. Basal cell carcinomas in mice overexpressing sonic hedgehog. Science 1997; 276:817-821. 44. Goodrich LV, Johnson RL, Milenkovic L et al. Conservation of the hedgehog/patched signaling pathway from flies to mice: Induction of a mouse patched gene by Hedgehog. Genes Dev 1996; 10:301-312. 45. Marigo V, Scott MP, Johnson RL et al. Conservation in hedgehog signaling: Induction of a chicken patched homolog by Sonic hedgehog in the developing limb. Development 1996b; 122:1225-1233. 46. Stone DM, Hynes M, Armanini M et al. The tumour-suppressor gene patched encodes a candidate receptor for Sonic hedgehog. Nature 1996; 384:129-134. 47. Ingham PW, Taylor AM, Nakano Y. Role of the Drosophila patched gene in positional signalling. Nature 1991; 353:184-187. 48. Hahn H, Christiansen J, Wicking C et al. A mammalian patched homolog is expressed in target tissues of sonic hedgehog and maps to a region associated with developmental abnormalities. J Biol Chem 1996; 271:12125-12128. 49. Kingsley DM. What do BMPs do in mammals? Clues from the mouse short-ear mutation. Trends Genet 1994; 10:16-21. 50. Graff, JM. Embryonic patterning: To BMP or not to BMP, that is the question. Cell 1997; 89:171-174. 51. Suzuki A, Kaneko E, Maeda J et al. Mesoderm induction by BMP-4 and -7 heterodimers. Biochem Biophys Res Commun 1997; 232:153-156. 52. Winnier G, Blessing M, Labosky PA et al. Bone morphogenetic protein-4 is required for mesoderm formation and patterning in the mouse. Genes Dev 1995; 9:2105-2116. 53. Heikinheimo K Stage-specific expression of Decapentaplegic-Vg-related genes 2, 4, and 6 (bone morphogenetic proteins 2, 4, and 6) during human tooth morphogenesis. J Dent Res 1994; 73:590-597. 54. Chen Y, Bei M, Woo I et al. Msx1 controls inductive signaling in mammalian tooth morphogenesis. Development 1996; 122:3035-3044. 55. Francis PH, Richardson MK, Brickell PM et al. Bone morphogenetic proteins and a signalling pathway that controls patterning in the developing chick limb. Development 1994; 120:209-218.

Early Molecular Events in Feather Morphogenesis

261

56. Francis-West PH, Robertson KE, Ede DA et al. Expression of genes encoding bone morphogenetic proteins and sonic hedgehog in talpid (ta3) limb buds: Their relationships in the signalling cascade involved in limb patterning. Dev Dyn 1995; 203:187-197. 57. Roberts DJ, Johnson RL, Burke AC et al. Sonic hedgehog is an endodermal signal inducing Bmp-4 and Hox genes during induction and regionalization of the chick hindgut. Development 1995; 121:3163-3174. 58. Staehling-Hampton K, Hoffmann FM, Baylies MK et al. Dpp induces mesodermal gene expression in Drosophila. Nature 1994; 372:783-786. 59. Kawakami Y, Ishikawa T, Shimabara M et al. BMP signaling during bone pattern determination in the developing limb. Development 1996; 122:3557-3566. 60. Zou H, Niswander L. Requirement for BMP signaling in interdigital apoptosis and scale formation. Science 1996; 272:738-741. 61. Parr BA, McMahon AP. Dorsalizing signal Wnt7a required for normal polarity of D-V and A-P axes of mouse limb. Nature 1995; 374:350-353. 62. McMahon AP, Moon RT. Ectopic expression of the proto-oncogene int-1 in Xenopus embryos leads to duplication of the embryonic axis. Cell 1989; 58:1075-1084. 63. Cui Y, Brown JD, Moon RT et al. Xwnt-8b: A maternally expressed Xenopus Wnt gene with a potential role in establishing the dorsoventral axis. Development 1995; 121:2177-2186. 64. Behrens J, von-Kries JP, Kuhl M et al. Functional interaction of β-catenin with the transcription factor LEF-1. Nature 1996; 382:638-642. 65. Tanda N, Ohuchi H, Yoshioka H et al. A chicken Wnt gene, Wnt-11, is involved in dermal development. Biochem Biophys Res Commun 1995; 211:123-129. 66. Davidson D. The function and evolution of Msx genes: Pointers and paradoxes. Trends Genet 1995; 11:405-411. 67. Maeda R, Kobayashi A, Sekine R et al. Xmsx-1 modifies mesodermal tissue pattern along dorsoventral axis in Xenopus laevis embryo. Development 1997; 124:2553-2560. 68. Thesleff I, Vaahtokari A, Partanen AM. Regulation of organogenesis. common molecular mechanisms regulating the development of teeth and other organs. Internat J Dev Biol 1995; 39:35-50. 69. Graham A, Koentges G, Lumsden A. Neural crest apoptosis and the establishment of craniofacial pattern: An honorable death. Mol Cell Neurosci 1996; 8:76-83. 70. Takahashi Y, Tonegawa A, Matsumoto K et al. BMP-4 mediates interacting signals between the neural tube and skin along the dorsal midline. Genes Cells 1996; 1:775-783. 71. Maas R, Chen YP, Bei M et al. The role of Msx genes in mammalian development. Ann N Y Acad Sci 1996; 785:171-181. 72. Liu YH, Kundu R, Wu L et al.Premature suture closure and ectopic cranial bone in mice expressing Msx2 transgenes in the developing skull.Proc Natl Acad Sci USA 1995; 92:6137-6141. 73. Chuong CM, Noveen AN, Jiang T-X et al. The homeobox gene Msx-2 is specifically expressed in epithelia destined to form skin appendages and is involved in the induction and growth of feather and hair. J Inv Dermatol 1995; 104:583. 74. Vogel A, Roberts-Clarke D, Niswander L. Effect of FGF on gene expression in chick limb bud cells in vivo and in vitro. Dev Biol 1995; 171:507-520. 75. Reardon W, Winter R, Rutland P et al. Mutations in the fibroblast growth factor receptor 2 gene cause Crouson syndrome. Nature Genet 1994; 8:98-103. 76. Noveen A, Jiang T-X, Ting-Berreth SA et al. Homeobox genes Msx-1 and Msx-2 are associated with induction and growth of skin appendages. J Invest Dermatol 1995a; 104:711-719. 77. McComb RB, Bowers GN, Rosen S. Alkaline Phosphatase. Plenum Press, New York. 1979. 78. Crawford K, Weissig H, Binnette F et al. Tissue-nonspecific alkaline phosphatase participates in the establishment and growth of feather germs in embryonic chick skin cultures. Dev Dyn 1995; 204:48-56. 79. Wessels NK. Morphology and proliferation during early feather development. Dev Biol 1965; 12:131-153. 80. Stuart ES, Moscona AA. Embryonic morphogenesis: Role of fibrous lattice in the development of feathers and feather patterns. Science 1967; 157:947-948.

262

Molecular Basis of Epithelial Appendage Morphogenesis

81. Goetinck PF, Sekellick MJ. Observations on collagen synthesis, lattice formation, and morphology of scaleless and normal embryonic skin. Dev Biol 1972; 28:636-648. 82. Marsh RG, Gallin WJ. Toxic effects of beta-aminopropionitrile treatment on developing chicken skin. J Exp Zool 1994; 268:381-389. 83. Goetinck PF, Carlone DL. Altered proteoglycan synthesis disrupts feather pattern formation in chick embryonic skin. Dev Biol 1988; 127:179-186. 84. Mauger A, Demarchez M, Herbage D et al. Immunofluorescent localization of collagen types I and III, and of fibronectin during feather morphogenesis in the chicken embryo. Dev Biol 1982; 94:93-105. 85. Chiquet-Ehrismann R, Mackie EJ, Pearson CA et al. Tenascin: An extracellular matrix protein involved in tissue interactions during fetal development and oncogenesis. Cell 1986; 47:131-139. 86. Jones FS, Burgoon MP, Hoffman S et al. A cDNA clone for cytotactin contains sequences similar to epidermal growth factor-like repeats and segments of fibronectin and fibrinogen. Proc Natl Acad Sci USA 1988; 85:2186-2190. 87. Erickson HP, Lightner VA. Hexabrachion protein (tenascin, cytotactin, brachionectin) in connective tissues, embryonic brain, and tumors. Adv Cell Biol 1988; 2:55-90. 88. Mackie EJ, Tucker RP, Halfter W et al. The distribution of tenascin coincides with pathways of neural crest cell migration. Development 1988; 102:237-250. 89. Hans-Henning E, Halfter W, Tucker RP. The distribution of fibronectin and tenascin along migratory pathways of the neural crest in the trunk of amphibian embryos. Development 1988; 103:743-756. 90. Jiang T-X, Chuong C-M. Mechanism of skin morphogenesis. I. Analyses with antibodies to adhesion molecules tenascin, N-CAM, and integrin. Dev Biol 1992; 150:82-98. 91. Noveen A, Jiang T-X, Chuong C-M. Protein kinase A and protein kinase C modulators have reciprocal effects on mesenchymal condensation during skin appendage morphogenesis. Dev Biol 1995b; 171:677-683. 92. Ruoslahti E. Integrins as signaling molecules and targets for tumor therapy. Kidney Int 1997; 51:1413-1417. 93. Chuong CM, Edelman GM. Expression of cell-adhesion molecules in embryonic induction. I. Morphogenesis of nestling feathers. J Cell Biol 1985; 101:1009-1026. 94. Fearon ER, Cho KR, Nigro JM et al. Identification of a chromosome 18q gene that is altered in colorectal cancers. Science 1990; 247:49-55. 95. Fearon ER, Pierceall WE. The deleted in colorectal cancer (DCC) gene: A candidate tumour suppressor gene encoding a cell surface protein with similarity to neural cell adhesion molecules. Cancer Surv 1995; 24:3-17. 96. Vielmetter J, Kayyem JF, Roman JM et al. Neogenin, an avian cell surface protein expressed during terminal neuronal differentiation, is closely related to the human tumor suppressor molecule deleted in colorectal cancer. J Cell Biol 1994; 127:2009-2020. 97. Chan SS, Zheng H, Su MW et al. UNC-40, a C. elegans homolog of DCC (Deleted in Colorectal Cancer), is required in motile cells responding to UNC-6 netrin cues. Cell 1996; 87:187-195. 98. Kolodziej PA, Timpe LC, Mitchell KJ et al. frazzled encodes a Drosophila member of the DCC immunoglobulin subfamily and is required for CNS and motor axon guidance. Cell 1996; 87:197-204. 99. Keino-Masu K, Masu M, Hinck L et al. Deleted in Colorectal Cancer (DCC) encodes a netrin receptor. Cell 1996; 87:175-185. 100. Kennedy TE, Serafini T, de la Torre JR et al. Netrins are diffusible chemotropic factors for commissural axons in the embryonic spinal cord. Cell 1994; 78:425-435. 101. Serafini T, Kennedy TE, Galko MJ et al. The netrins define a family of axon outgrowthpromoting proteins homologous to C. elegans UNC-6. Cell 1994; 78:409-424. 102. Colamarino SA, Tessier-Lavigne M. The axonal chemoattractant netrin-1 is also a chemorepellent for trochlear motor axons. Cell 1995; 81:621-629.

Early Molecular Events in Feather Morphogenesis

263

103. Wadsworth WG, Bhatt H, Hedgecock EM. Neuroglia and pioneer neurons express UNC-6 to provide global and local netrin cues for guiding migrations in C. elegans. Neuron 1996; 16:35-46. 104. Chuong CM, Jiang T-X, Yin E et al. cDCC (Chicken homologue to a gene deleted in colorectal carcinoma) is an epithelial adhesion molecule expressed in the basal cells and involved in epithelial-mesenchymal interaction. Dev Biol 1994; 164:383-397. 105. Huttenlocher A, Sandborg RR, Horwitz AF. Adhesion in cell migration. Curr Opin Cell Biol 1995; 7:697-706. 106. Gille J, Swerlick RA. Integrins: Role in cell adhesion and communication. Ann N Y Acad Sci 1996; 797:93-106. 107. Moses HL, Arteaga CL, Alexandrow MG et al. TGF beta regulation of cell proliferation. Princess Takamatsu Symp 1994; 24:250-263. 108. Chiquet-Ehrismann R, Kalla P, Pearson CA. Participation of tenascin and transforming growth factor-beta in reciprocal epithelial-mesenchymal interactions of MCF7 cells and fibroblasts. Cancer Res 1989; 49:4322-4325. 109. Nakashima M, Nagasawa H, Yamada Y et al. Regulatory role of transforming growth factor-beta, bone morphogenetic protein-2, and protein-4 on gene expression of extracellular matrix proteins and differentiation of dental pulp cells. Dev Biol 1994; 162:18-28. 110. Jakowlew SB, Ciment G, Tuan RS et al. Expression of transforming growth factor-beta 2 and beta 3 mRNAs and proteins in the developing chicken embryo. Differentiation 1994; 55:105-118. 111. Ting-Berreth SA, Chuong C-M. Local delivery of TGF β2 can substitute for placode epithelium to induce mesenchymal condensation during skin appendage morphogenesis. Dev Biol 1996b; 179:347-359. 112. Massague J, Weis-Garcia F. Serine/threonine kinase receptors: Mediators of transforming growth factor beta family signals. Cancer Surv 1996; 27:41-64. 113. Iseki S, Araga A, Ohuchi H et al. Sonic hedgehog is expressed in epithelial cells during development of whisker, hair, and tooth. Biochem Biophys Res Commun 1996; 218:688-693. 114. Noveen A, Jiang TX, Chuong CM. cAMP, an activator of protein kinase A, suppresses the expression of sonic hedgehog. Biochem Biophys Res Commun 1996; 219:180-185. 115. Desbiens X, Queva C, Jaffredo T et al. The relationship between cell proliferation and the transcription of the nuclear oncogenes c-Myc, c-myb and c-Ets-1 during feather morphogenesis in the chick embryo. Development 1991; 111: 699-713. 116. Turque N, Buttice G, Beuscart A et al.Hydrocortisone modulates the expression of c-Ets-1 and 72 kDa type IV collagenase in chicken dermis during early feather morphogenesis. Int J Dev Biol 1997; 41:103-109. 117. Gurdon JB, Harger P, Mitchell A et al. Activin signalling and response to a morphogen gradient. Nature 1994; 371:487-492. 118. Held LI. Models for embryonic periodicity. Karger, New York 1992; 16-24.

CHAPTER 14

Signaling Loops in the Reciprocal Epithelial-Mesenchymal Interactions of Mammalian Tooth Development Yi-Ping Chen and Richard Maas

O

rganogenesis is a complex process that results from sequential and reciprocal cell and tissue interactions. Tooth development, which shares similarities with the development of other embryonic organs, is characterized by a series of reciprocal epithelial-mesenchymal interactions that result in the differentiation and organization of the interacting tissues. Therefore, mammalian tooth development provides an excellent model for studying epithelial-mesenchymal interactions. The recent combination of molecular biology and classical experimental embryology has begun to elucidate the molecular mechanisms of such interactions in tooth morphogenesis. In situ hybridization and immunochemical analysis have revealed the expression of growth factors such as BMPs, FGFs, and other signaling molecules, and of transcription factors such as homeobox containing genes in different developmental phases and tissue compartments of the developing tooth germ. The function of growth factors as inductive signaling molecules in epithelial-mesenchymal interactions during early tooth development has been demonstrated by their ability to substitute for a tissue requirement in the induction of gene expression and morphologic changes in an adjacent tissue. Similarly, the importance of transcription factors such as Msx-1, Msx-2, Dlx1, Dlx2, Lef1 and Pax9 in tooth development has also been demonstrated, by elimination of the gene products using gene targeting. Mice deficient for any of these genes exhibit a severe tooth phenotype. The striking correlation of growth factor gene expression with that of transcription factors in interacting epithelial and mesenchymal tissues during organogenesis suggests a general model whereby growth factors, which can diffuse between interacting tissues, both regulate and are regulated by transcription factors. Such a model postulates recursively utilized inductive loops to account for the sequential and reciprocal signaling events that operate in organogenesis. Molecular analysis of tooth development in Msx-1 deficient mice supports the hypothesis that one function of homeobox genes is to regulate the expression of growth factors, since Msx genes function to permit inductive signaling to occur back and forth between tissue layers.

Introduction Vertebrate organs form through sequential and reciprocal interactions between apposed epithelial and mesenchymal tissues. These processes involve a series of inductive and permissive tissue interactions (also termed secondary induction) which eventually lead to Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

266

Molecular Basis of Epithelial Appendage Morphogenesis

cell differentiation and organ formation. It has been demonstrated that each tissue layer is required for the growth and differentiation of the other. Neither epithelium nor mesenchyme can grow and differentiate if cultured alone.1-3 In the case of those organs that form via epithelial-mesenchymal interactions, the initiation of vertebrate organ formation first occurs as a local thickening of the epithelium, which subsequently invaginates into the subjacent mesenchymal cells. This process is accompanied by the condensation of mesenchymal cells. Further morphogenesis of developing organs involves rapid and complex growth of epithelium, such as branching within the mesenchyme and/or folding around the condensed and differentiating mesenchymal cells.4 Classical tissue recombination experiments have demonstrated that the developmental potential of organ-specific morphogenesis resides either in the epithelium or mesenchyme, depending on the organ and its developmental stage. Although studies on the molecular basis of vertebrate appendage development have focused on limb morphogenesis (reviewed in refs. 5-6), substantial information on the molecular mechanisms of tooth development is rapidly accumulating (reviewed in refs. 4 and 7). Tooth development is characterized by a series of sequential and reciprocal epithelial-mesenchymal interactions which regulate the initiation, shaping and terminal differentiation of dentition. Murine tooth development has proven to be a powerful experimental system for studying inductive tissue interactions and regulation of organogenesis. The possibility of in vitro organ culture, easy access to cytological and molecular studies, and the clear delineation of epithelial and mesenchymal components for tissue recombination make the developing tooth a highly attractive experimental model for elucidating the molecular regulation of inductive tissue interactions.8 Similar to the development of other organs such as kidney, lung, mammary gland and hair follicle, tooth formation is also traditionally divided into three partly overlapping phases: initiation, morphogenesis and differentiation.9 In the mouse embryo, initiation of molar tooth development begins morphologically at embryonic day 11.5 (E11.5) with a visible thickening of dental epithelium to form the dental lamina. The dental lamina subsequently invaginates into the underlying neural crestderived dental mesenchyme to form the early and later bud stage at E12.5 and E13.5, respectively. Meanwhile, the surrounding dental mesenchyme proliferates and condenses. Thereafter, during the cap stage (E14.5), the epithelial bud becomes convoluted to form a cap-like structure, which marks the first definitive stage of tooth morphogenesis. The cap stage is followed by the bell stage (E16.5), which is characterized by more convoluted epithelial structures. Characteristic features of these later stages include the development of the enamel epithelium and formation of the enamel knot, a putative organizing center proposed to provide positional information for tooth morphogenesis and regulate the growth of tooth cusps.10 Postnatally, these epithelial-mesenchymal interactions culminate in the differentiation of the epithelium into highly specialized enamel-secreting ameloblasts and of the mesenchyme into dentin-secreting odontoblasts and alveolar bone. For a more detailed discussion of tooth morphogenesis, see chapter 9. Over the last decade, the application of molecular biology to problems in development has provided insight into the molecular mechanisms of vertebrate organogenesis. A large body of evidence has indicated that the epithelial-mesenchymal interactions which govern early tooth morphogenesis are accompanied by the expression of numerous genes in both tissue layers. These include growth factors and their receptors, transcription factors, and extracellular matrix and membrane proteins. Members of several growth factor families, including transforming growth factor beta (TGF-β), bone morphogenetic proteins (BMP), fibroblast growth factors (FGF), epidermal growth factor (EGF) and Sonic hedgehog (Shh) have been detected in the developing tooth germ (reviewed in ref. 4).

Signaling Loops in Mammalian Tooth Development

267

General Models for Signaling Loops in Tissue Interactions Several lines of evidence demonstrate that growth factors function in tissue interactions by acting as inductive signals. Their expression in different tissue layers at different stages is consistent with a major function for growth factors as potential inductive signals in the sequential and reciprocal epithelial-mesenchymal interactions that occur during organogenesis. A general model, for which there is considerable experimental support, is presented in Figure 14.1 to show how growth factors and transcription factors can function recursively during inductive tissue interactions. In this model, a specific growth factor (GF1) or set of growth factors (GF1a,b,c...), are expressed in the initial inductive tissue, tissue A, and their gene products diffuse into the interacting tissue, tissue B where cognate receptors are expressed. Activation of these receptors leads to the expression of specific transcription factors (TFa,b,c...) which are necessary for the expression of a second set of growth factors (GF2a,b,c...) in the responding tissue, tissue B. Although these factors may exert tissue autonomous functions, they can also then feed back upon the original inductor, tissue A, to form a signaling loop during organogenesis. In the context of tooth development, BMP-4 fulfills the criteria of factor GF1, while Msx-1 fulfills that for TF1. Other BMPs, FGFs and potentially Shh and Wnts are candidates for other GFs, while Dlxs, Pax gene products, members of the paired homeodomain family and HMG box members such as Lef1 are candidates for the TFs. Two other features are implicit in the model proposed in Figure 14.1. First, if the only function of the transcription factors TF1a,b,c..., TF2a,b,c... in organogenesis were the reciprocal exchange of signals back and forth between the interacting tissues, there would be no basis for the successive differentiation of the tissues. Thus, although not specifically indicated in Figure 14.1, it must be understood that in addition to regulating growth factor gene expression, the transcription factors must also regulate the expression of structural genes at each developmental step. Second, it should be noted that the transcription factors may be necessary but not sufficient to activate the expression of various downstream genes, implying that specific combinations of transcription factors are required. The necessary condition is usually inferred from an absence of gene expression in a loss of function mutant. The sufficient condition is more difficult to establish. In some cases, however, it can be shown in organ culture experiments that the induction of transcription factor gene expression fails to induce the expression of a growth factor gene presumed to reside genetically downstream. Although unproven, we consider it likely that specific combinations of transcription factors must therefore interact to achieve positive regulation of the various growth factor genes. It is possible that combinations of different signals are used to define the specific “code” of transcription factors that in turn specifies which growth factor genes will be reciprocally expressed. The potential complexity of this code could engender some of the specificity inherent in the differentiation of organs along different fates, and the progression of this “code” at each developmental step would dictate individual organ fates. This review focuses specifically on the nature of the inductive signals which are exchanged between the dental epithelium and mesenchyme, and on one particular signaling loop controlled by Msx homeobox genes during early tooth morphogenesis which exemplifies the model presented in Figure 14.1.

Expression of Growth Factors Is Dynamic During Early Tooth Development A paradigm for growth factors as diffusible inductive signals between tissue layers was first established and elucidated for mesoderm induction in Xenopus embryos. Members of the TGF-β superfamily and FGF family have been shown to be critical signals in the specification of mesoderm by endoderm.11 These factors are expressed correctly in place and time,

268

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 14.1. A sequential and reciprocal model for functions of growth factors and transcription factors in the epithelial-mesenchymal interactions during vertebrate organogenesis. See text for details. GF: growth factor; TF: transcription factor.

and can substitute for the endoderm in mesoderm induction.12-13 Involvement of growth factors in mammalian organogenesis has also been demonstrated, including limb bud (reviewed in ref. 6), somite,14-16 and neural tube,17-18 suggesting that the same growth factors are repeatedly used as inductive signals in different morphogenetic tissue contexts during embryogenesis. Most evidence that growth factors act as inductive signals in epithelial-mesenchymal interaction has been obtained from studies of expression patterns of genes and proteins. Therefore, the expression of some important growth factors believed to function in tooth morphogenesis will be first discussed.

BMPs Bone Morphogenetic Proteins (BMPs) were originally defined by their ability to initiate bone induction in vertebrate tissues in vitro (reviewed in ref. 19). BMPs have been shown to play critical roles in mesoderm induction as well as in organ formation by acting as inductive signals.20-26 These proteins function as homodimers through binding to their receptors. However, in some cases, different BMPs may act in concert by forming BMP heterodimers (e.g., BMP-2/7 and BMP-4/7), further increasing potential complexity. Bmp-2, Bmp-4 and Bmp-7 are all expressed in dynamic patterns in the developing molar tooth germ. BMP-2 and BMP-4 share 95% amino acid sequence identity and belong to the same BMP subgroup. The vertebrate Bmp-2 and Bmp-4 genes are most closely related (about 75%) to the prototypical Decapentaplegic (Dpp) gene in Drosophila. Interestingly, it is evident that Dpp is also involved in tissue-tissue interactions in Drosophila. The role of Dpp as an inductive signal is exemplified by its ability to specify midgut constriction27-28 and to function in the induction of visceral and cardiac mesoderm.29 It was recently shown that Dpp can act at a distance by long-range diffusion, suggesting Dpp is a morphogen in Drosophila organogenesis.30-31 These facts suggest that the genetic pathways involving Dpp and BMPs may be conserved between invertebrates and vertebrates, as observed for the con-

Signaling Loops in Mammalian Tooth Development

269

served role of Hox genes in the specification of anterioposterior axial identity. During early murine tooth morphogenesis, Bmp-2 expression first appears mesially in the molar dental epithelium at late E11.32 The transcripts of Bmp-2 remain localized to the dental epithelium at E13.5, with expression confined to the middle of the epithelium, the prospective enamel knot.10 At the cap stage (E14.5), Bmp-2 is only expressed in the enamel knot region but not in the dental mesenchyme. Similar to Bmp-2, Bmp-7 expression is confined to the prospective enamel knot region at the bud stage and to the enamel knot at the cap stage. In contrast to the expression of Bmp-2 and Bmp-7, Bmp-4 transcripts are first detected in the thickened presumptive dental epithelium at E11. Subsequently, the expression domain expands to the underlying dental mesenchyme at E12, and then completely shifts to the condensing dental mesenchyme at E13.25 At the cap stage, the transcripts of Bmp-4 stay in the dental mesenchyme and expand to the area of the enamel knot of the dental epithelium.10 In addition, Bmp-3, Bmp-5 and Bmp-6 were not detected in the developing tooth germ at the bud and cap stages.10 It has been previously demonstrated by in vitro tissue recombination that tooth developmental potential shifts between tissue layers during early tooth morphogenesis.33-34 Prior to E12.5, the dental epithelium possesses tooth developmental potential, as shown by its ability to elicit tooth formation when combined with neural crest derived second branchial arch mesenchyme, while the reciprocal combination fails.33 After E12.5, this tooth forming inductive potential shifts to the dental mesenchyme, which now can induce tooth formation when recombined with second branchial arch epithelium. Interestingly, the changes of Bmp-4 expression pattern in the developing tooth germ coincide temporally with the shift in tooth developmental potential from epithelium to mesenchyme.25 These observations strongly suggest that the shift in Bmp-4 expression from dental epithelium to mesenchyme accounts for the transfer in inductive potential from dental epithelium to the mesenchyme, and that BMP-4 may represent one component of the tooth developmental potential. Due to its ability to induce gene expression and morphologic changes in the dental mesenchyme, BMP-4 has been proposed to be a critical inductive signal in early murine tooth development.25,35 Thus, considering the model proposed in Figure 14.1, BMP-4 may represent one of the initial signaling molecules (GF1) in Tissue A as well as, coincidentally, one of the signaling molecules (GF2) in Tissue B.

FGFs Evidence from the studies of mesoderm induction show that members of the FGF family also play critical roles in the regional specification of mesoderm, and in some cases have synergistic effects with TGF-β family growth factors in Xenopus embryos.13,36-37 Furthermore, a role for FGFs in vertebrate organogenesis is being established. For example, FGFs are required for limb initiation and outgrowth, as well as the maintenance of signaling loops during limb morphogenesis.38-42 Recent data from in situ hybridization and immunostaining experiments suggest a potential important role of FGFs in murine tooth morphogenesis. At least four FGF members, including FGF-3,-4,-7 and -8, have been shown to be expressed in the developing mouse tooth germ. Of these, Fgf-8 is the only one which is detected in the thickened dental epithelium at E11.5.43 The expression domain of Fgf-8 is restricted to the tooth forming region, providing an early marker for the presumptive dental epithelium (Chen and Maas, unpublished data). This early and restricted expression of Fgf-8 suggests that FGF-8 may also represent one of the early signaling molecules (GF1 in Fig. 14.1) expressed in the dental lamina during the initiation of tooth development. In contrast to Fgf-8, transcripts of Fgf-3 appear in the dental mesenchyme at the bud stage and are extensively expressed in the dental papilla until the bell stage.4,44 Restriction of Fgf-3 transcripts in the dental mesenchyme suggests its role in mesenchymal cell proliferation

270

Molecular Basis of Epithelial Appendage Morphogenesis

and in the determination of cell fate. Furthermore, it may represent an inductive signal from the dental mesenchyme to the epithelium during the progression of the bud to the cap stage. However, in Fgf-3 knockout mice, no obvious dental defects were reported,45 suggesting possible functional redundance between FGF-3 and other FGFs. Transcripts of Fgf-7, which encodes KGF (keratinocyte growth factor) are expressed in the dental mesenchyme, while its receptor KGFR was found in adjacent dental epithelial structures.46 These observations suggest that FGF-7 may represent another signaling molecule that exerts a developmental function on the dental epithelium. Lastly, Fgf-4 is specifically expressed in the epithelial enamel knot at E14.5 of molar tooth development. No Fgf-4 transcripts prior to the cap stage have been detected in the developing tooth germ.10

Other Signaling Molecules The role of epidermal growth factor (EGF) in tooth morphogenesis was suggested by the presence of its transcripts in both the epithelial and mesenchymal components of the developing mandibular arches at E9 and E10.47 EGF transcripts also colocalize with EGF receptor (Egfr) to the dental epithelium and mesenchyme at the bud stage of incisor and molar tooth development.48 The requirement for EGF in odontogenesis has been demonstrated by antisense abrogation experiments, where inhibition of EGF expression by antisense oligodeoxynucleotides blocks tooth development.48-49 It was recently suggested that EGF may interact with retinoic acid in controlling the pattern of the dentition by affecting dental lamina patterning.50 TGF-α, which belongs to the EGF family and shares the same receptor (EGFR) with EGF is also expressed in the developing tooth germ. TGF-α transcripts localize to the dental epithelium at the cap stage, while immunostaining studies revealed the localization of TGF-α protein to both dental epithelium and mesenchyme.51 These observations indicate that autocrine and paracrine effects of TGF-α during odontogenesis are likely. Mutation of TGF-α in mice does not cause odontogenic defects.52-53 However, functional redundance between EGF and TGF-α may account for these observations. The TGF-β superfamily comprises about 20 members, including TGF-β1-β5. The transcripts or proteins of TGF-β1-3 have been detected in the mammalian developing tooth germ.54-56 TGF-β1 mRNA is initially localized to the dental epithelium at the bud stage, and remains specifically in the epithelial structures through the cap and bell stages.57 This restricted expression of TGF-β1 in the dental epithelium appears to be regulated by signals from dental mesenchyme.57 TGF-β2 proteins were found to localize in the dental lamina at the initiation stage and were found in both epithelium and mesenchyme at the bud stage.56 Interestingly, the transcripts of TGF-β3 were detected only in the dental mesenchyme at the bud stage,54 while TGF-β3 proteins were found to be present in the dental epithelium and basement membrane.56 Since several TGF-β members are present in the developing tooth germ, a functional redundance between these TGF-βs may exist. Both type I and type II TGF-β receptors are expressed in the dental epithelium and mesenchyme at the bud and cap stages,58,59 suggesting their role in the signal transduction of TGF-βs in early tooth morphogenesis. Another TGF-β superfamily member which plays an important role in embryonic mesoderm induction is the activin βA gene.11,60 This gene is also expressed in dental mesenchyme at E13.5, suggesting a potential role in the progression from the bud to cap stage. Indeed, knockout mice lacking the activin βA gene exhibit lack of incisors and immature molar tooth buds.61 Recent work from several laboratories has revealed that Sonic hedgehog (Shh), a critical signaling molecule participating in pattern formation of several organs,17,62 is expressed in the developing tooth germ. Although Shh is expressed only at low levels in the molar dental epithelium at the initiation stage,63,64 the expression of Shh in the epithelial enamel knot has been well documented.10,64 Actually, the expression of Shh in the developing molar

Signaling Loops in Mammalian Tooth Development

271

tooth germ is now employed as a molecular marker for the enamel knot structure and for the determination of tooth developmental stage. The coexpression of Shh and Bmp genes in the ectodermal or adjacent mesenchymal structures bears striking similarity to patterning events in Drosophila imaginal discs, where hedgehog (hh) signaling is responsible for the induction and maintenance of Dpp expression.65-67 Ectopic expression of Shh in the avian developing limb and gut induces ectopic expression of Bmp-2 and Bmp-4, respectively.68,69 The use of this same signaling pathway in both flies and vertebrates suggests the evolutionary conservation of this signaling cascade. The recently cloned mouse homolog of the patched gene encodes a transmembrane protein which can function as a receptor for Shh and can mediate Shh function.70-72 It has been shown that patched is upregulated by Shh in the developing limb,73 neural tube70 and tooth (Chen and Maas, unpublished data). In the early tooth germ, patched expression is asymmetrically localized in the dental mesenchyme at E11.5 (Chen and Maas, unpublished data), and remains in the dental mesenchyme until at least the bud stage.70

Expression of Transcription Factors Is Associated with Inductive Processes Transcription factors are DNA-binding proteins that directly control the activity of other genes. In situ hybridization experiments in past years have revealed the expression of several important transcription factor genes in the mammalian developing tooth germ. Among these are Msx-1, Msx-2, Lef1, Dlx1, Dlx2, Dlx3, Pax9 and Egr1. These genes exhibit unique expression patterns that overlap with those growth factors in the developing tooth germ, suggesting a potential relationship between the two classes of gene products in inductive interactions.

Msx The vertebrate Msx gene family was originally identified on the basis of homeobox sequence homology to the Drosophila msh gene (muscle segment homeobox gene).74,75 The mammalian Msx gene family consists of 3 physically unlinked members, named Msx-1, Msx-2 and Msx-3. Msx-3 thus far has only been found to be expressed in dorsal neural tube, resembling the expression pattern of the prototypical Drosophila msh gene.76 In developing vertebrate embryos, Msx-1 and Msx-2 are widely expressed in many organs, including neural crest, limb bud, heart, hair follicles and teeth. Of particular interest is that these two genes are expressed at almost all sites of epithelial-mesenchymal interactions (reviewed in ref. 77). The unique expression pattern of Msx-1 and Msx-2 strongly suggests a role for these two genes in the formation of organs governed by epithelial-mesenchymal interactions. Msx-1 and Msx-2 are expressed in the developing tooth germ with distinct temporal and spatial patterns which correlate with discrete morphologic steps in tooth development.78-80 At E10.5, Msx-2 expression is first detected in the mesenchyme beneath the site of dental placode formation, representing the earliest molecular marker for odontogenesis.80 Slightly later, at E11.5, both Msx-1 and Msx-2 are coexpressed in presumptive dental mesenchyme. Thereafter, while the expression of both Msx-1 and Msx-2 persists in the dental mesenchyme, a component of Msx-2 expression shifts to the epithelium, where it becomes restricted to the enamel knot. Thus, Msx-1 and Msx-2 are expressed in both overlapping and distinct patterns in the early developing tooth germ. In addition to their expression patterns, increasing evidence also points to the involvement of Msx genes in epithelial-mesenchymal interactions. Msx expression in various mesenchymal tissues can be induced by epithelial contact.81-83 This epithelial induction of Msx gene expression in the mesenchyme is also positionally dependent. Heterospecific grafts between chick and mouse limb and facial primordia have shown that Msx-1 expression is

272

Molecular Basis of Epithelial Appendage Morphogenesis

activated when facial mesenchyme is grafted into the limb, but not the converse.84 These results indicate that expression of Msx genes in the grafted mesenchyme is in accord with the fate of its new host enviroment.84 The epithelial-mesenchymal signaling systems are partially but not completely interchangeable. This epithelial induction of Msx expression is also demonstrated in the developing tooth germ by in vitro tissue recombination. Dental mesenchymal expression of Msx-1 and Msx-2 can be induced by dental epithelium until the E16 bell and E13 bud stages, respectively.85 Similarly, in the mouse embryonic limb, mesenchymal expression of Msx-1 depends upon the presence of ectoderm.86 These results strongly suggest that signals from adjacent tissue layers are required for the expression of Msx genes. The importance of Msx genes in epithelial-mesenchymal interactions during organogenesis was demonstrated by genetic experiments in which the function of Msx-1 and Msx-2 has been eliminated. Msx-1 or Msx-2 deficient mice show defects in organs such as teeth which form by epithelial-mesenchymal interactions. Msx-1 deficient mice exhibit cleft palate and an arrest of tooth development at the bud stage.87 In contrast, Msx-2 knockout mice exhibit normal early tooth development. However, detailed analysis of teeth from Msx-2 deficient mice reveals defects in both incisor and molar tooth development, including a reduction in the amount of stellate reticulum and abnormal patterning of the future cusp regions (Satokata et al, unpublished data). Since Msx-1 and Msx-2 are frequently expressed in overlapping domains in many developing organs, and their protein sequence differs by only 2 amino acids within the DNA binding homeodomain, functional redundancy may exist between Msx-1 and Msx-2 during embryogenesis. Consistent with this hypothesis, mice deficient for both Msx-1 and Msx-2 manifest severe defects in numerous organs (Satokata et al, unpublished data). In Msx-1-Msx-2 double mutants, tooth development is arrested at the initiation stage, indicating that Msx-1 and Msx-2 gene function is required in the dental mesenchyme as early as E11.5.

Lef1 Lymphoid enhancer factor 1 (Lef1) is a cell type specific architectural protein that belongs to the HMG box family (reviewed in ref. 88). Lef1 protein does not directly activate transcription, but instead induces a sharp bend in the DNA helix, facilitating the ability of other transcription factors to activate transcription.89 The expression of Lef1 is restricted to lymphocytes in the adult mouse, and in the neural crest, mesencephalon, tooth germs, hair follicles and other sites during embryogenesis.89-93 In the developing tooth germ, while Lef1 transcripts are first detected in the thickened dental epithelium at E11, expression shifts to the subjacent dental mesenchyme at E12. At E13, Lef1 is expressed both in the condensed dental mesenchyme and in the immediate adjacent basal cells of the dental epithelium. This expression pattern remains throughout the subsequent cap and bell stages.93 These sites of expression overlap those identified for Msx-1 and Msx-2. Loss of Lef1 function results in mice lacking teeth, mammary gland, hair and whiskers, which form through sequential epithelial-mesenchymal interactions.91 Thus, Lef1 may play a general regulatory role in epithelial-mesenchymal interactions during organogenesis. In the context of tooth formation, Lef1 mutant mice exhibit normal initiation but an arrest of tooth development at the bud stage, similar to that in Msx-1 mutant mice.87,91 Further studies using classical tissue recombination between Lef1 mutant and wild type embryos revealed that Lef1 function is required only transiently in the dental epithelium to induce morphogenesis in the mesenchyme.93 Since the expression of Lef1 precisely overlaps with Msx-1 and Msx-2 in the developing tooth germ and, since tooth development arrests at essentially the same stage in both Msx-1 and Lef1 mutant mice, it is possible that Msx-1 and Lef1 reside within a common genetic pathway.

Signaling Loops in Mammalian Tooth Development

273

Other Transcription Factors The mammalian Distal-less (Dlx) gene family consists of at least 6 distinct genes whose sequences show homology to the Drosophila distal-less gene. The transcripts of Dlx1, 2, 3 and 5 are present in the murine developing tooth germ by RT-PCR.94 Dlx1 is initially expressed in the dental mesenchyme at E11.5, and its expression remains there until at least the cap stage.95 Dlx2 is expressed in the dental epithelium at E12 to E14, with weak expression in the adjacent dental mesenchyme.96 Thus, the expression pattern of Dlx1 and Dlx2 coincides with mesenchymal Msx-1 and epithelial Msx-2 expression in the developing tooth germ. Dlx3 expression is specifically restricted in the dental mesenchyme at the bud and cap stages.96 Although neither Dlx1 nor Dlx2 knockout mice show a tooth phenotype, Dlx1Dlx2 double mutant mice exhibit an arrest in maxillary molar tooth development at the early bud stage.97 The coexpression of Dlx and Msx genes and growth factors in the developing tooth germ suggests that Dlx genes may participate in tooth morphogenesis through genetic pathways similar to those involving Msx and growth factors. Pax9, a member of paired domain family genes, is also expressed in the dental mesenchyme at the bud stage.98 Elimination of Pax9 function in mice results in arrest of tooth development at the bud stage.98 It will be very interesting to know whether Pax9 resides in the same genetic pathway with Msx genes in controlling tooth development.

Growth Factors as Signals Mediating Reciprocal Epithelial-Mesenchymal Interactions Although little is known about the determination of tooth initiation sites, it is generally accepted that the first inductive signals come from the thickened dental epithelium at the initiation stage during tooth morphogenesis. The epithelial signals act instructively on the underlying dental mesenchyme and elicit a sequential and reciprocal signaling cascade. This early signaling process results in induction of various molecules in the subjacent dental mesenchyme, and morphologically in the condensation of surrounding dental mesenchymal cells. The inductive function of dental epithelial signals is demonstrated by in vitro tissue recombination where the dental epithelium and mesenchyme of early tooth germs are enzymatically separated and then reassociated on Nuclepore filters in Trowell type organ culture (reviewed in ref. 99). After 24-48 h culture, the expression of specific mesenchymal molecules is analyzed by in situ hybridization and immunostaining. It has been shown that the early dental epithelium is able to induce the expression of Msx-1, Msx-2, Egr-1, syndecan-1, tenascin as well as Bmp-4 in the dental mesenchyme.25,35,85,100,101 As controls, no gene expression could be detected in the mesenchyme cultured alone. The induction of gene expression is accompanied by the condensation of adjacent mesenchymal cells. Therefore, the in vitro recombinants essentially recapitulate the early events of tooth morphogenesis in vivo. Egr-1 (Early growth response gene-1) encodes a protein containing three DNAbinding zinc finger sequences and is specifically expressed in the dental mesenchyme at the bud and cap stages.101,102 It has been suggested that Egr-1 may be involved in tissue interactions and in the determination of cell fate in the developing tooth germ.101 The cell surface heparan sulfate proteoglycan syndecan-1 and the extracellular matrix glycoprotein tenascin are cell adhesion molecules and are specifically localized to the condensing mesenchyme during the early process of tooth development.103 Syndecan-1 from the embryonic tooth mesenchyme binds to tenascin,104 pointing to their role in mesenchymal cell condensation.4 In addition, epithelial signals also stimulate cell proliferation in the adjacent dental mesenchyme. This induced cell proliferation is closely associated with induction of syndecan-1, suggesting that the two events may be related.100 Since syndecan-1 contains heparan and chondroitin sulfate side chains, it may participate in growth control by binding to growth factors, most likely FGFs, and thereby modulate the availability and activity of growth factors.

274

Molecular Basis of Epithelial Appendage Morphogenesis

An elegant experiment to test whether epithelial signals are diffusible growth factors was designed by Thesleff and colleagues, in which a growth factor-soaked bead replaces the dental epithelium in the in vitro recombination cultures described above (reviewed in ref. 7). In this system, beads soaked with growth factors are placed on the top of, or adjacent to, the dental mesenchyme. Induction of gene expression in the mesenchyme is assayed after a period of time in culture. Since Bmp-2 and Bmp-4 are expressed in the thickened dental epithelium at the initiation stage, these are attractive molecules for the initial inductive signals.25 Indeed, both BMP-2 and BMP-4 releasing beads are able to substitute for some effects of the presumptive dental epithelium in the induction of gene expression and morphologic changes in the dental mesenchyme. BMP-4 releasing beads induce expression of transcription factors Msx-1, Msx-2, Lef1 and Egr-1 in dental mesenchyme.25,35,93 Furthermore, BMP-4 beads also induce translucent zones in the adjacent mesenchyme, mimicking the effect of the dental epithelium.25 Interestingly, BMP-4 has also been shown to induce the expression of Bmp-4 itself in the mesenchyme.25,35 The inductive effect of BMP-4 on the mesenchyme has been suggested to be mediated through a diffusion mechanism, although a homeogenetic induction was not excluded.25 As discussed below, the induced BMP-4 in the dental mesenchyme may in turn represent one of the inductive signals which exert their function on the dental epithelium for its further development and differentiation, and thus a signaling loop is formed during the epithelial-mesenchymal interactions. Despite the dramatic inductive effects of BMPs on tooth morphogenesis, BMPs alone are not sufficient to substitute for all endogenous signaling functions of the dental epithelium. Neither expression of syndecan-1 and tenascin nor cell proliferation can be induced by BMP releasing beads in the dental mesenchyme.25 Since Fgf-8 is also expressed in the thickened dental epithelium of E11.5 murine tooth germ, FGFs may represent another component of epithelial signaling. This hypothesis is supported by evidence that FGFs, including FGF-1,-2 and-4, can substitute for the dental epithelium in the induction of syndecan1 and tenascin as well as cell proliferation in the dental mesenchyme.25,35,99,105 In addition, FGF-4 inhibits apoptosis in the dental mesenchyme.106 On the other hand, similar to BMPs, FGFs are also potent inducers of Msx-1 expression in the dental mesenchyme,35 and induce translucent zones in the adjacent mesenchyme.99 Since none of these tested FGFs are expressed in the dental epithelium at the initiation stage, they may not be the endogenous initial epithelial signals. The inductive effect of these FGFs may mimic the activity of FGF-8.35 The subsequent expression of several growth factors in the dental mesenchyme at the bud stage may play a role in the commitment of mesenchymal cells to the odontogenic lineage as well as in the regulation of tooth-specific epithelial morphogenesis. Mesenchymally expressed growth factors thus potentially represent subsequent reciprocal signals which can exert their inductive functions on the epithelium. Mesenchymally derived signals are required for gene expression in the dental epithelium, as exemplified by the induction of TGF-β1 expression in the dental epithelium by signals from the dental mesenchyme.57 Among the putative signaling molecules, BMP-4, FGF-3, FGF-7, activin βA and TGF-β3 all are expressed in the dental mesenchyme at the bud and following stages. These mesenchymal signals, as evident for BMP-4, may be involved in the progression of tooth development from the bud to the cap stage and in the induction of the enamel knot.35 Moreover, they may also control subsequent tooth epithelial morphogenesis, as demonstrated by tissue recombination studies where dental papilla instructs tooth formation when recombined with epithelium from other sources.3 Several studies have shown that the transcripts of Fgf-4, Bmp-2, Bmp-4, Bmp-7 as well as Shh are localized to the enamel knot at the cap stage.10,64,105 Interestingly, all these signaling molecules are found in the apical ectodermal ridge (AER) and/or the zone of polarizing activity (ZPA) of the developing vertebrate limb, where the AER is required for outgrowth

Signaling Loops in Mammalian Tooth Development

275

and the ZPA for anterioposterior patterning (reviewed in ref. 6). In addition, there exist positive signaling feedback loops operating between AER and ZPA to coordinately regulate the expression of signaling molecules in each tissue.6,39 These observations indicate that a similar signaling mechanism may be employed by these two structurally divergent organs during organogenesis. In support of this hypothesis, it was demonstrated that grafts of mouse tooth germ into the anterior domain of the chick developing limb bud induce digit duplication in the host limb, mimicking the effect of the ZPA of the limb bud.64 These findings suggest that the epithelially derived enamel knot is a signaling center which may function in later tooth patterning and differentiation.10,99 This discovery provides a new insight that instead of dental mesenchyme alone, the dental epithelium is also involved in the regulation of tooth shape.10,105 It is conceivable that signaling from the enamel knot can also act back on the dental mesenchyme to stimulate cell growth and differentiation, as evidenced by the effect of FGF-4 in the stimulation of cell proliferation and the inhibition of apoptosis in the dental mesenchyme.105-106

MSX Genes Regulate the Expression of Inductive Signals The coexpression of homeobox-containing genes with growth factors at many sites during embryogenesis suggests a close link between them in the regulation of gene expression. Although it is clear that growth factors regulate homeobox gene expression, several lines of evidence indicate the regulation of growth factors by homeobox genes. For instance, Dpp expression is directly regulated in visceral mesoderm by the Ultrabithorax (Ubx) homeoprotein in Drosophila embryos.27 The homeobox genes Msx-1 and Msx-2 have been implicated in the epithelial-mesenchymal interactions mediated by growth factors. The precise role of Msx genes in regulating such interactions during organogenesis has been further exploited by using Msx-1 mutant embryos. The Msx-1 mutant tooth germ was used as a model system, because Msx-1 mutant mice exhibit an arrest in molar tooth development at the bud stage (E13.5). This arrest is accompanied by a marked reduction in the degree of mesenchymal condensation.87 Since Msx-1 is expressed only in the dental mesenchyme during early odontogenesis and since several growth factors are coexpressed with Msx-1 in the dental mesenchyme at the bud stage, Msx-1 may function in the dental mesenchyme for the progression of tooth development beyond the bud stage by controlling the expression of growth factors. Consistent with this model are studies that E11.5 Msx-1 mutant tooth germs in in vitro organ culture supplemented with 10% fetal calf serum can develop to the cap stage, the stage that E11.5 wild type tooth germs usually reach in similar conditions (Fig. 14.2). These observations suggest that diffusible factors can bypass Msx-1 function in the dental mesenchyme. Growth factors which are coexpressed with Msx-1 in the dental mesenchyme at E13.5, including BMP-4, FGF-3, TGF-β3 and activin βA, represent potential downstream target genes regulated by the Msx-1 gene product in the dental mesenchyme. Bmp-4, whose expression pattern shifts with the transfer of tooth developmental potential from the dental epithelium to mesenchyme at the time when tooth development is arrested in Msx-1 mutant mice, appears to be a particularly compelling Msx-1 downstream target gene. In situ hybridization and RT-PCR analyses demonstrate that Bmp-4 expression in the dental epithelium in the E11.5 Msx-1 mutant tooth germ is preserved, but the expression in the E13.5 Msx-1 mutant dental mesenchyme is dramatically reduced compared with that of wild type dental mesenchyme.35 These observations were further confirmed by immunochemical analysis using an antibody against BMP-4 and BMP-2. Moreover, by using an in vitro bead experiment in mesenchymal explants, it was shown that Msx-1 is required for the induction of Bmp-4 by BMP-4 itself in the dental mesenchyme.35 Based on these observations, Msx-1 expression not only responds to induction by BMP-4, but is also required for the expression

276

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 14.2. Partial rescue of Msx-1 mutant tooth phenotype by diffusible factors. (A) Tooth development reaches the cap stage in an E14.5 wild type embryo. (B) Tooth development is arrested at the bud stage in an E14.5 Msx-1 mutant embryo. (C) Tooth structure at the cap stage is observed in an E11.5 wild type tooth germ after 5 days in culture in vitro with 10% FCS. (D) An E11.5 Msx-1 mutant tooth germ developed beyond the bud stage to the cap stage after 5 days in culture in vitro with 10% FCS. Without FCS, mutant tooth germs remain arrested at the bud stage.

of Bmp-4 in the dental mesenchyme. In addition, Lef1 expression is also markedly reduced in Msx-1 mutant dental mesenchyme at E13.5. These results place mesenchymally expressed Bmp-4 and Lef1 downstream of Msx-1 during early tooth development. The observations that Lef1 can be induced by BMP-4 in the dental mesenchyme35,93indicate that reduction of BMP-4 may account for the downregulation of Lef1 expression in the Msx-1 mutant dental mesenchyme. All the results described above make it possible to order these genes into a genetic pathway during early tooth morphogenesis (Fig. 14.3). Lastly, in vitro rescue experiments in which E13.5 Msx-1 mutant tooth germ cultured with exogenous BMP-4 can develop to the cap stage demonstrate that mesenchymally expressed BMP-4 is one of the inductive signals acting upon the dental epithelium. This idea is supported by the fact that ALK-3, a Type I BMP-receptor serine-threonine kinase which preferentially binds BMP-4, is expressed at E12.5 in the dental epithelium.107-108 Exogenous BMP-4 can bypass the function of Msx-1 in the dental mesenchyme at the bud stage and partly rescue the tooth phenotype in Msx-1 mutants (Chen et al, 1996). Thus, Msx-1 indeed regulates expression of inductive signals such as BMP-4 which are required for further morphogenesis of dental epithelium (Fig. 14.3). It would be very interesting to examine the expression of other signaling molecules in the Msx-1 mutant dental mesenchyme. This regulation of growth factors by Msx genes in early tooth development may be conserved in other developing organs that form via sequential epithelial-mesenchymal interactions. The observations that Msx genes also regulate Bmp expression in the developing limb support this hypothesis (Chen and Maas, unpublished data).

Signaling Loops in Mammalian Tooth Development

277

Fig. 14.3. A model integrating Msx-1, Bmp-4 and Lef1 into a genetic pathway in early tooth development. Signaling loops are formed both between two tissue layers and within a tissue layer. The arrows are not necessarily intended to indicate direct interactions, only an epistasis relationship. Reprinted with permission of Company of Biologists Ltd. from Chen YP et al, Development 1996; 122:3035-3044.

In addition to its role in the regulation of growth factor expression, Msx genes may be involved in cell condensation and the expression of cell adhesion molecules.109 In the Msx-1 mutant dental mesenchyme, lesser degrees of cellular condensation are observed.87 The expression of syndecan-1, but not tenascin, is significantly reduced in E13.5 Msx-1 mutant dental mesenchyme.35 Preservation of tenascin expression in the Msx-1 mutant dental mesenchyme argues against the possibility that the arrest of tooth development is caused by a general defect in neural crest derived mesenchyme. The reduction of syndecan-1 expression may account for the lesser degrees of mesenchymal cell condensation in the Msx-1 mutant tooth germ. It was further shown that although FGFs can induce syndecan-1 expression in the dental mesenchyme, the induction of syndecan-1 by FGFs is mediated by Msx gene products.35 Although evidence presented here to support the signaling loop model (Fig. 14.1) derives from studies of murine tooth development, this model should have general implications for vertebrate organogenesis. In summary, recent advances have revealed important insights into the nature of inductive signaling loops that operate in the context of epithelialmesenchymal interaction during vertebrate organogenesis.

Acknowledgment The research discussed here from the authors’ laboratory is supported by an NIH grant (RO1DE11697) to RM, and an NIH NRSA (DE05671) and NSF grant (IBN 96-03801) to YPC.

278

Molecular Basis of Epithelial Appendage Morphogenesis

References 1. Grobstein C. Induction interaction in the development of the mouse metanephros. J Exp Zool 1955; 130:319-340. 2. Kollar E.J, Baird GR. The influence of the dental papilla on the development of tooth shape in embryonic mouse tooth germ. J Embryol Exp Morphol 1996; 21:131-148. 3. Kollar EJ, Baird GR. Tissue interactions in embryonic mouse tooth germs: II. The inductive role of the dental papilla. J Embryol Exp Morphol 1970; 24:173-186. 4. Thesleff I, Vaahtokari A, Partanen A. Regulation of organogenesis. Common molecular mechanisms regulating the development of teeth and other organs. Int J Dev Biol 1995; 39:35-50. 5. Tickle C, Eichele G. Vertebrate limb development. Annu Rev Cell Biol 1994; 10:121-152. 6. Tickle C. Vertebrate limb development. Sem Cell Dev Biol 1996; 7:137-143. 7. Thesleff I, Vaahtokari A, Vainio S et al. Molecular mechanisms of cell and tissue interactions during early tooth development. Anat Rec 1996; 245: 151-161. 8. Maas R, Bei M. The genetic control of early tooth development. Crit Rev Oral Biol Med 1997; 8:4-39. 9. Kollar E.J, Lumsden AGS. Tooth morphogenesis: The role of the innervation during induction and pattern formation. J Biol Buccale 1979; 7:49-60. 10. Vaahtokari A, Åberg T, Jernvall J et al. The enamel knot as a signaling center in the developing mouse tooth. Mech Dev 1996; 54:39-43. 11. Jessell TM, Melton DA. Diffusible factors in vertebrate embryonic induction. Cell 1992; 68:257-270. 12. Kessler DS, Melton DA. Vertebrate embryonic induction: Mesodermal and neural patterning. Science 1994; 266:596-604. 13. Slack JWW. Inducing factors in Xenopus early embryos. Curr Biol 1994; 4:116-126. 14. Fan C-M, Tessier-Lavigne M. Patterning of mammalian somites by surface ectoderm and notochord: Evidence for sclerotome induction by a hedgehog homolog. Cell 1994; 79:1175-1186. 15. Munsterberg AE, Kitajewski J, Bumcrot DA et al. Combinatorial signaling by sonic hedgehog and Wnt family members induces myogenic bHLH gene expression in the somites. Genes Dev 1995; 9:2911-2922. 16. Pourquié O, Fan C-M, Coltey M et al. Lateral and axial signals involved in avian somite patterning: A role for BMP-4. Cell 1996; 84:461-471. 17. Roelink H, Porter JA, Chiang C et al. Floor plate and motor neuron induction by different concentrations of amino-terminal cleavage product of sonic hedgehog autoproteolysis. Cell 1995; 81:445-455. 18. Liem KF Jr, Tremml G, Roelink HR et al. Dorsal differentiation of neural plate cells induced by BMP-mediated signals from epidermal ectoderm. Cell 1995; 82:969-979. 19. Kingsley DM. What do BMPs do in mammals? Clues from the mouse short-ear mutation. Trends Genet 1994; 10:16-21. 20. Hemmati-Brivanlou A, Thomsen G. Ventral mesodermal patterning in Xenopus embryos: Expression patterns and activities of BMP-2 and BMP-4. Dev Genet 1995; 17:78-89. 21. Lyons KM, Jones CM, Hogan BLM. The DVR gene family in embryonic development. Trends Genet 1991; 7:408-412. 22. Heikinheimo K. Stage-specific expression of Decapentaplegic-Vg-related genes 2, 4, and 6 (bone morphogenetic protein 2, 4 and 6) during human tooth morphogenesis. J Dent Res 1994; 73:590-597. 23. Blessing M, Nanney LB, King LE et al. Transgenic mice as model to study the role of TGF-β-related molecules in hair follicles. Genes Dev 1993; 7:204-215. 24. Francis PH, Richardson MK, Brichell PM et al. Bone morphogenetic proteins and signaling pathway that controls patterning in the developing chick limb. Development 1994; 120:209-218. 25. Vainio S, Karavanova I, Jowett A et al. Identification of BMP-4 as a signal mediating secondary induction between epithelial and mesenchymal tissues during tooth development. Cell 1993; 75:45-58.

Signaling Loops in Mammalian Tooth Development

279

26. Hogan BLM. Bone morphogenetic proteins: Multifunctional regulators of vertebrate development. Genes Dev 1996; 10:1580-1594. 27. Capovilla M, Brandt M, Botas G. Direct regulation of Decapentaplegic by Ultrabithorax and its role in midgut morphogenesis. Cell 1994; 76:461-476. 28. Bienz M. Homeotic genes and positional signaling in the Drosophila viscera. Trends Genet 1994; 10:22-26. 29. Frasch M. Induction of visceral and cardiac mesoderm by ectodermal Dpp in the early Drosophila embryo. Nature 1995; 374:464-467. 30. Lecuit T, Brook WJ, Ng M et al. Two distinct mechanisms for long-range patterning by Decapentaplegic in the Drosophila wing. Nature 1996; 381:387-393. 31. Nellen D, Burke R, Struhl G et al. Direct and long-range action of a DPP morphogen gradient. Cell 1996; 85:357-368. 32. Turecková J, Sahberg C, Åberg T et al. Comparison of expression of the msx-1, msx2, BMP-2 and BMP-4 genes in the mouse upper diastemal and molar primordia. Int J Dev Biol 1995; 39:459-468. 33. Mina M, Kollar EJ. The induction of odontogenesis in non-dental mesenchyme combined with early murine mandibular arch epithelium. Arch Oral Biol 1987; 32:123-127. 34. Lumsden AGS. Spatial organization of the epithelium and the role of neural crest cells in the initiation of the mammalian tooth germ. Development Supplement 1988; 103:155-169. 35. Chen YP, Bei M. Msx-1 controls inductive signaling in mammalian tooth morphogenesis. Development 1996; 122:3035-3044. 36. Kimmelman D, Kirschner M. Synergistic induction of mesoderm by FGF and TGF-β and the identification of an mRNA coding for FGF in the early Xenopus embryo. Cell 1987; 51:868-877. 37. Reilly KM, Melton DA. The role of short-range and long-range signaling in mesoderm induction and patterning during Xenopus development. Sem Cell Dev Biol 1996; 7:77-85. 38. Niswander L, Tickle C, Vogel A et al. FGF-4 replaces the apical ectodermal ridge and directs outgrowth and patterning of the limb. Cell 1993; 75:579-587. 39. Niswander L, Jeffrey S, Martin GR et al. A positive feedback loop coordinates growth and patterning in the vertebrate limb. Nature 1994; 371:609-612. 40. Cohn MJ, Izpisúa-Belmonte JC, Abud H et al. Fibroblast growth factors induce additional limb development from the flank of chick embryos. Cell 1995; 80:739-746. 41. Crossley PH, Minowada G, MacArthur C et al. Roles for FGF-8 in the induction, initiation, and maintenance of chick limb development. Cell. 1996; 84:127-136. 42. Fallon JE, Lopez A, Ros MA et al. FGF-2: Apical ectodermal ridge growth signal for chick limb development. Science 1994; 264:104-107. 43. Heikinheimo M, Lawshe A, Shackleford GM et al. FGF-8 expression in the post-gastrulation mouse suggests role in the development of the face, limb and central nervous system. Mech Dev 1994; 48:129-138. 44. Wilkinson DG, Bhatt S, McMahon AP et al. Expression pattern of the FGF-related protooncogene int-2 suggests multiple roles in fetal development. Development 1989; 105:131-136. 45. Mansour SL, Goddard JM, Capecchi MR et al. Mice homozygous for a targeted disruption of the proto-oncogene int-2 have developmental defects in the tail and inner ear. Development 1993; 117:13-28. 46. Finch PW, Cunha GR, Rubin JS et al. Pattern of keratinocyte growth factor and keratinocyte growth factor receptor expression during mouse fetal development suggests a role in mediating morphogenetic mesenchymal-epithelial interactions. Dev Dyn 1995; 203:223-240. 47. Kronmiller JE, Upholt WB. Expression of epidermal growth factor mRNA in the developing mouse mandibular process. Arch Oral Biol 1991; 36:405-410. 48. Shum L, Sakakura Y, Bringas P Jr et al. EGF abrogation-induced fusilli-form dysmorphogennesis of Meckel’s cartilage during embryonic mouse mandibular morphogenesis in vitro. Development 1993; 118:903-917. 49. Kronmiller JE, Upholt WB, Kollar EJ et al. EGF antisense oligodeoxynucleotides block murine odontogenesis in vitro. Dev Biol 1991; 147:485-488.

280

Molecular Basis of Epithelial Appendage Morphogenesis

50. Kronmiller JE. Spatial distribution of epidermal growth-factor transcripts and effects of exogenous epidermal growth factor on the pattern of the mouse dental lamina. Arch Oral Biol 1995; 40:137-143. 51. Huang L, Solursh M, Sandra A. The role of transforming growth factor alpha in rat craniofacial development and chondrogenesis. J Anat 1996; 189:73-86. 52. Luetteke NC, Qiu TH, Peiffer RL et al. TGF-α Deficiency results in hair follicle and eye abnormalities in targeted and waved-1 mice. Cell 1993; 73:263-278. 53. Mann GB, Fowler KJ, Gabriel A et al. Mice with a null mutation of the TGF-α gene have abnormal skin architecture, wavy hair, and curly whiskers and often develop corneal inflammation. Cell 1993; 73:249-261. 54. Pelton R, Dickinson ME. In situ hybridization analysis of TGF-β3 RNA expression during mouse development: Comparative studies with TGF-β 1 and β 2. Development 1990; 110:609-620. 55. Pelton R, Saxena B. Immunohistochemical localization of TGF-β1, TGF-β2 and TGF-β3 in the mouse embryo: Expression patterns suggest multiple roles during embryonic development. J Cell Biol 1991; 115:1091-1105. 56. Chai Y, Mah A, Crohin C et al. Specific transforming growth factor-β subtypes regulate embryonic mouse Meckel’s cartilage and tooth development. Dev Biol 1994; 162:85-103. 57. Vaahtokari A, Vainio S, Thesleff I et al. Associations between transforming growth factor β1 RNA expression and epithelial-mesenchymal interactions during tooth morphogenesis. Development 1991; 113:985-994. 58. Wang Y, Sizeland A, Wang XF et al. Restricted expression of type-II TGF-β receptor in murine embryonic development suggests a central role in tissue modeling and CNS patterning. Mech Dev 1995; 52:275-289. 59. Iseki S, Osumi-Yamashita N, Miyazono K et al. Localization of transforming growth factor-beta type I and type II receptors in mouse development. Exp Cell Res 1995; 219:339-347. 60. Thomsen GT, Woolf T, Whitmen M et al. Activins are expressed early in Xenopus embryogenesis and can induce axial mesoderm and anterior structures. Cell 1990; 63:485-493. 61. Matzuk MM, Kumar TR, Vassalli A et al. Functional analysis of activins during mammalian development. Nature 1995; 375:354-356. 62. Riddle RD, Johnson RJ, Laufer E et al. Sonic hegdehog mediates the polarizing activity of ZPA. Cell 1993; 75:1401-1416. 63. Bitgood MJ, McMahon AP. Hedgehog and BMP genes are coexpressed at many diverse sites of cell-cell interaction in the mouse embryo. Dev Biol 1995; 172:126-138. 64. Koyama E, Yamaai T, Iseki S et al. Polarizing activity, sonic hedgehog, and tooth development in embryonic and postnatal mouse. Dev Dyn 1996; 206:59-72. 65. Basler K, Struhl K. Compartment boundaries and the control of Drosophila limb pattern by hedgehog protein. Nature 1994; 368:208-214. 66. Ma C, Zhou Y, Beachy PA et al. The segment polarity gene hedgehog is required for progression of the morphogenetic furrow in the developing Drosophila eye. Cell 1993; 75:913-926. 67. Tabata T, Kornberg TB. Hedgehog is a signaling protein with a key role in patterning Drosophila imaginal discs. Cell 1994; 76:89-102. 68. Laufer E, Nelson CE, Johnson RL et al. Sonic hedgehog and Fgf-4 act through a signaling cascade and feedback loop to integrate growth and patterning on the developing limb bud. Cell 1994; 79:993-1003. 69. Roberts DJ, Johnson RL, Burke AC et al. Sonic hedgehog is an endodermal signal inducing Bmp-4 and Hox genes during induction and regionalization of the chick hindgut. Development 1995; 121:3163-3174. 70. Goodrich LV, Johnson R.L, Milenkovic L et al. Conservation of the hedgehog/patched signaling pathway from flies to mice: Induction of a mouse patched gene by Hedgehog. Genes Dev 1996; 10:301-312. 71. Stone DM, Hynes M, Armanini M et al. The tumour-suppressor gene patched encodes a candidate receptor for Sonic hedgehog. Nature 1996; 384:129-134.

Signaling Loops in Mammalian Tooth Development

281

72. Marigo V, Davey RA, Zuo Y et al. Biochemical evidence that Patched is the Hedgehog receptor. Nature 1996; 384:176-179. 73. Marigo V, Scott M, Johnson RL et al. Conservation in hedgehog signaling: Induction of a chicken patched homolog by Sonic hedgehog in the developing limb. Development 1996; 122:1225-1233. 74. Hill RE, Jones PF, Rees AR et al. A new family of mouse homeo box containing genes: Molecular structure, chromosomal location, and developmental expression of Hox-7.1. Genes Dev 1989; 3:26-37. 75. Robert B, Sassoon D, Jacq B et al. Hox-7, a mouse homeobox with a novel pattern of expression during embryogenesis. EMBO J 1989; 8:91-100. 76. Shimeld S, McKay IJ, Sharpe PT et al. The murine homeobox gene Msx-3 shows highly restricted expression in the developing neural tube. Mech Dev 1996; 55:201-210. 77. Davidson D. The function and evolution of Msx genes: Pointers and paradoxes. Trends Genet 1995; 11:405-411. 78. MacKenzie A, Leeming GL, Jowett AK et al. The homeobox gene Hox7.1 has specific regional and temporal expression patterns during early murine craniofacial embryogenesis, especially tooth development in vivo and in vitro. Development 1991; 111:269-285. 79. MacKenzie A, Ferguson MWJ, Sharpe PT. Hox-7 expression during murine craniofacial development. Development 1991; 113:601-611. 80. MacKenzie A, Ferguson MWJ, Sharpe PT. Expression patterns of the homeobox gene, Hox8, in the mouse embryo suggest a role in specifying tooth initiation and shape. Development 1992; 115:403-420. 81. Robert B, Lyons G, Sinandl BK et al. The apical ectodermal ridge regulates Hox-7 and Hox-8 gene expression in developing chick limb buds. Genes Dev 1991; 5:2363-2374. 82. Takahashi Y, Bontoux M, Le Dourin NM. Epithelial-mesenchymal interactions are critical for Quox7 expression and membrane bone differentiation in the neural crest derived mandibular mesenchyme. EMBO J 1991; 10:2387-2393. 83. Pavlova A, Boutin E, Cunha G et al. Msx-1 (Hox-7) in the adult mouse uterus: Cellular interactions underlying regulation of expression. Development 1994; 120:335-346. 84. Brown JM, Wedden SE, Millburn GH et al. Experimental analysis of the control of expression of the homeobox-gene Msx-1 in the developing limb and face. Development 1993; 119:41-48. 85. Jowett AK, Vainio S, Furguson MW et al. Epithelial-mesenchymal interactions are required for msx1 and msx2 gene expression in the developing murine molar tooth. Development 1993; 117:461-470. 86. Wang Y, Sassoon D. Ectoderm-mesenchyme and mesenchyme-mesenchyme interactions regulate Msx-1 expression and cellular differentiation in the murine limb bud. Dev Biol 1995; 68:374-382. 87. Satokata I, Maas R. Msx-1 deficient mice exhibit cleft palate and abnormalities of craniofacial and tooth development. Nature Genet 1994; 6:348-356. 88. Grosschedl R, Giese K, Pagel J. HMG domain proteins: Architectural elements in the assembly of nucleoprotein structures. Trends Genet 1994; 10:94-100. 89. Travis A, Amsterdam A, Belanger C et al. LEF-1, a gene encoding a lymphoid-specific protein with an HMG domain, regulates T-cell receptor α enhancer function. Genes Dev 1991; 5:880-894. 90. Oosterwegel M, van de Wetering J, Timmerman J et al. Differential expression of the HMG boxfactors TCF-1 and LEF-1 during murine embryogenesis. Development 1993; 118:439-448. 91. van Genderen C, Okamura RM, Farinas I et al. Development of several organs that required inductive epithelial-mesenchymal interactions is impaired in LEF-1-deficient mice. Genes Dev 1994; 8:2691-2703. 92. Zhou P, Byrne C, Jacobs J et al. Lymphoid enhancer factor 1 directs hair follicle patterning and epithelial cell fate. Genes Dev 1995; 9:700-713. 93. Kratochwil K, Dull M, Farinas I et al. Lef1 expression is activated by BMP-4 and regulates inductive tissue interactions in tooth and hair development. Genes Dev 1996; 10:1382-1394.

282

Molecular Basis of Epithelial Appendage Morphogenesis

94. Weiss K.M, Bollekens FH, Ruddle FH et al. Distal-less and other homeobox genes in the development of the dentition. J Exp Zool 1994; 270:273-284. 95. Dollé P, Price M, Duboule D. Expression of the murine Dlx-1 homeobox gene during facial, ocular and limb development. Differentiation 1992; 49:93-99. 96. Robinson GW, Mahon KA. Differential and overlapping expression domains of Dlx-2 and Dlx-3 suggest distict roles for Distal-less homeobox genes in craniofacial development. Mech Dev 1994; 48:199-215. 97. Thomas BL et al. Role of Dlx1 and Dlx2 genes in patterning of murine dentition. Development 1997; 124:4811-4818. 98. Peters H et al. Pax9 deficient mice lack pharyngeal pouches derivatives and teeth, and exhibit craniofacial and limb abnormalitites. Genes and Dev (in press). 99. Thesleff I, Sahlberg C. Growth factors as inductive signals regulating tooth morphogenesis. Sem Cell Dev Biol 1996; 7:185-193. 100. Vainio S, Thesleff I. Coordinated induction of cell proliferation and syndecan expression in dental mesenchyme by epithelium: Evidence for diffusible signals. Dev Dyn 1992; 194:105-117. 101. Karavanova I, Vainio S, Thesleff I. Transient and recurrent expression of the Egr-1 gene in epithelial and mesenchymal cells during tooth morphogenesis suggests involvement in tissue interactions and in determination of cell fate. Mech Dev 1992; 39:41-50. 102. McMahon AP, Champion JE, McMahon JA et al. Developmental expression of the putative transcription factor Egr-1 suggests that Egr-1 and c-fos are coregulated in some tissues. Development 1990; 108:281-287. 103. Vainio S, Thesleff I. Sequential induction of syndecan, tenascin and cell proliferation associated with mesenchymal cell condensation during early tooth development. Differentiation 1992; 50:97-105. 104. Salmivirta M, Elenius K, Vainio S et al. Syndecan from embryonic tooth mesenchyme binds tenascin. J Biol Chem 1991; 266:7733-7739. 105. Jernvall J, Kettunen P, Karavanova I et al. Evidence for the role of the enamel knot as a control center in mammalian tooth cusp formation: Non-dividing cells express growth stimulating Fgf-4 gene. Int J Dev Biol 1994; 38:463-469. 106. Vaahtokari A, Åberg T, Thesleff I. Apoptosis in the developing tooth: Association with an embryonic signaling center and suppression by EGF and FGF-4. Development 1996 122:121-129. 107. Dewulf N, Verschueren K, Lonnoy O et al. Distinct spatial and temporal expression patterns of two type I receptors for bone morphogenetic proteins during mouse embryogenesis. Endocrinology 1995; 136:2652-2663. 108. ten Dijke P, Yamashita H, Sampath TK et al. Identification of type I receptors for osteogenic protein-1 and bone morphogenic protein-4. J Biol Chem. 1994; 269:16985-16988. 109. Hall BK, Miyake T. Divide, accumulate, differentiate: Cell condensation in skeletal development revisited. Int J Dev Biol 1995; 39:881-893.

CHAPTER 15

Topobiology of the Hair Follicle: Adhesion Molecules as Morphoregulatory Signals During Hair Follicle Morphogenesis Sven Müller-Röver and Ralf Paus

Introduction

O

ne fundamental question in hair follicle (HF) development is: How can a tiny cluster of seemingly uniform epithelial cells, which associates with a small cluster of apparently homogeneous mesenchymal cells, give rise to a complex mini-organ that serves as an enormously productive fiber factory—the hair follicle? While there is ever-increasing interest in the role of key growth factors/cytokines and transcription factors in the control of HF development1-3 (chapter 6), much less attention has been paid to the role that adhesion molecules may play in this context.4-6 This is regrettable, since it is now clear that adhesion molecules are crucially involved in translating the one-dimensional genetic code into a threedimensional tissue architecture and that adhesion-dependent form, placement and migration of cells are among the basic driving forces of morphogenesis.7-11 Since substantial evidence already points to a major role of adhesion molecules in closely related developmental systems, such as tooth and feather development (see chapters 9, 13), it is advisable to consider adhesion molecules as an eminent force in hair morphogenesis, as well. In order to illuminate the complex and probably crucial functions of adhesion molecules in HF development, and to stimulate more systematic research efforts in this important, yet neglected area of hair biology, we shall first delineate some basic principles concerning the role of adhesion molecules in developmental systems and shall briefly introduce some major protagonists that may be particularly relevant to hair research. This is followed by a short account of Edelman's “Morphoregulator Hypothesis”; and a definition of key questions in HF “topobiology”. We then review the currently available, limited evidence for a possible involvement of selected cadherins, integrins and members of the immunoglobulin superfamily in HF development, before closing with some working hypotheses on the role of these adhesion molecules in HF morphogenesis so as to guide future studies in this field.

Principles of Topobiology in Morphogenesis In order to better understand the role that adhesion molecules may play in HF morphogenesis,4-6 it is useful to first consider basic principles by which cell-cell and cell-matrix adhesion, as well as cell migration, act to shape morphogenesis. We will therefore briefly Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

284

Molecular Basis of Epithelial Appendage Morphogenesis

review the function of major adhesion molecules and reflect on how the expression of adhesion molecules may be controlled and coordinated.

Control of Morphogenesis by Cell Adhesion and Migration Morphogenesis is driven by the basic phenomena of cell proliferation, migration and differentiation as well as by programmed cell death, the establishment of cell polarity, and the joining of cooperating cell collectives into functional tissue units. All these are, in turn, controlled—at least to a significant extent—by adhesion forces. During embryonic development, coherent groups of cells segregate, aggregate, and obtain positional information and polarity on the basis of their adhesion properties, which create differential surface tension forces and alter cell shape.10,11 While it is undisputed that cell state (e.g., cell cycle state, genetically determined differentiation program) and cell lineage are critical elements in pattern formation during the morphogenesis of tissues and organs, and though it is clear that pattern formation can be brought about by a large diversity of interdependent mechanisms9 (see chapter 18), positional information on the basis of a cell’s distinct adhesion properties is surely of central importance.7 In fact, even the survival of individual cells during and after development appears to be critically controlled by adhesion forces that dictate cell shape.12,13 Highlighting the key role of positional information, Edelman8 has coined the term “topobiology” for the field that investigates the role of “place-dependent molecular interactions of cell surfaces or with substrates that result in changes in cell regulation”. According to the thermodynamic model of cell interactions in multicellular structures,14-18 cells rearrange themselves in the most thermodynamically stable pattern. Thereby, they exist in a state of equilibrium that is only disturbed by changes in gene expression which alter the cell surface expression of adhesion molecules, thus forcing the cells to move until a new, thermodynamically stable equilibrium has been attained.18 For example, when two cell populations differ only in the amount of cell surface expression of one particular adhesion molecule, they segregate to approach a sphere-within-a-sphere configuration; the cells with higher binding capacities (population A) adhere tightly to each other, and get surrounded by the cells with lower binding capacities (population B), thus assuming a fundamental, recurrent theme in development—that of “cortex” and “medulla” formation.18 Folding and invagination of epithelial sheets (e.g., during neurulation or HF development) operates on the principle that cells unevenly enlarge their surface on one side (e.g., the basal cell pole) so that the existing adhesion milieu creates bending forces which lead to folding or invagination of the epithelial sheet.11 Specific cell receptors for extracellular matrix molecules also guide the migratory movements of individual cells and entire cell collectives. These molecules permit, e.g., the movement of mesenchymal cells and neurons and allow the spatial separation of epithelial sheets.7,10,11 In addition, adhesion molecules join cell collectives to functional units by coordinating the gene expression, cell cycle and/or apoptosis of large cell collectives. Finally, adhesion molecules provide permeability barriers and impart mechanical strength to epithelial sheets, and serve as the basis of “positional memory” during tissue regeneration and cyclic tissue remodeling.7,10,11 Thus, the list of functional properties demonstrated for adhesion molecules has grown steadily over the past years to extend far beyond the control of mechanical adhesion. For example, adhesion molecules are now recognized to determine cell surface tension, to bind to extracellular matrix and/or the internal cytoskeleton, to modulate the activity of growth factor receptors, and/or to transduce signals that alter gene expression, thus modulating such basic biological phenomena as cell proliferation, apoptosis, migration, differentiation, and secretory activities.11,19,20

Topobiology of the Hair Follicle

285

Adhesion Molecules Potentially Relevant in Hair Research Several classes of adhesion molecules are distinguished, e.g., selectins, lectins, glycosyltransferases, cadherins, CAMs of the immunoglobulin superfamily and integrins, which display homophilic or heterophilic interactions (i.e., they bind to each other, or to selected binding partners, respectively).19,20 In this chapter, we shall focus only on the latter three families of adhesion molecules, since several pieces of evidence support that these are potentially relevant to hair biology. Cadherins The cadherins are a family of calcium-dependent cell-adhesion molecules that display homophilic as well as heterophilic binding. Over 30 different cadherins have already been identified, which are further subdivided into different subclasses with distinct binding specificities and tissue distribution. Cadherins are of major importance for assorting cells into tissues21-26 and for establishing cell polarity,27-32 and selected cadherins are critical factors protecting cells against apoptosis.31,33-35 Three major regions of cadherins can be distinguished: the extracellular region mediates specific adhesion, the transmembrane connects the intra- and extracellular region, and the intracellular region is linked to cytoskeleton by proteins that are termed catenins. These, in turn, bind to a diverse number of intracellular molecules such as the cytoskeletal proteins actin and α-actinin, the transcription factor Lef1 and members of the ErbB family of receptor tyrosine kinases, e.g., the epidermal growth factor receptor.21,22,36 Here, we focus on the classical cadherins, E- and P-cadherin (Ecad, Pcad), which are the best studied as well as the predominant cadherins found in epithelial cells. Ecad and Pcad expression has been demonstrated in the hair follicle (Table 15.1). Ecad is crucial to the generation and maintenance of epithelial sheets during morphogenesis and later tissue homeostasis.37 The disruption of Ecad-mediated adhesion can lead to a dedifferentiation of epithelial cells to a fibroblast-like morphology, while exogenous introduction of Ecad cDNA forces nonepithelial cell lines to form epithelial-like monolayers with a polarized phenotype.38,39 Ecad can induce junctional complexes such as the adherens junctions40-42 and inhibit cell proliferation.42 Furthermore, Ecad expression may be a crucial factor in determining the intraepithelial location of Langerhans cells43 and γδ-TCR+ lymphocytes.44,45 Particular relevant in our context is that Ecad may modulate the activity of the transcription factor Lef1 via beta-catenin46 which, in turn, may regulate the expression of Ecad47 (see below). In addition, it has been suggested that epidermal stem cells express relatively high levels of gamma-catenin and lower levels of Ecad and betacatenin than other basal layer epidermal keratinocytes (KC) destined to undergo terminal differentiation.48 Placental or P-cadherin (Pcad) is highly expressed in the murine placenta during pregnancy, but also in several other tissues including the epidermis.49 Quantitative differences in P-cadherin expression dramatically influence cell sorting in vitro and contribute to threedimensional pattern formation, such as the generation of “cortex” and “medulla” structures from precursor cell populations which differ only in their amount of Pcad surface expression.18 In skin biology, Pcad may be important for segregating the proliferating KC of the basal epidermal layer from the suprabasal layers.50 Cell-adhesion molecules (CAMs) of the immunoglobulin superfamily This family of calcium-independent cell-adhesion molecules is very heterogeneous, and numerous different functional roles have been described for the over 70 members of the immunoglobulin superfamily. Some CAMs operate primarily as signal transducing receptors (such as the IL-1 or PDGF-receptors), while others also provide mechanochemical links between cells (such as NCAM). The best known members of this family include the

Molecular Basis of Epithelial Appendage Morphogenesis

286

Table 15.1. E- and P-cadherin expression during murine and human skin and hair follicle morphogenesis Species Skin Type

E-Cadherin

P-Cadherin

Ref. 122

mouse

fetal upper • basal and intermediate lip skin epidermal layer • ORS • outer HM and proximal IRS

• basal epidermal layer • ORS • inner HM

mouse

neonatal back skin

• all epidermal layers 123,124 including hair pegs • progressive downregulation on all epidermal layers except basal layer; strong IR on developing ORS, inner HM, keratogenic region of precortical HM

• all epidermal layers • progressive downregulation on developing inner HM and slightly on developing IRS and keratogenic region of precortical HM; strong expression on myoepithelial cells of SG, ORS, outer HM

human fetal head, • all epidermal cells including • basal cells trunk and basal cells (weaker), spinous • ORS, inner HM, SG (hair breast skin cells and granular cells peg + bulbous hair peg) • central portion of hair pegs • colocalization with Ki-67 and follicles and ORS (proliferation marker) • ORS, IRS, outer HM, SG (hair peg + bulbous hair peg)

50

human fetal thigh and trunk skin

125

• all borders of intermediate cells (pregerm) • weak expression on hair germ (in contrast to strong suprabasal staining) • moderate expression on hair peg, weaker expression on HM cells close to the DP • all cells of bulbous hair peg, except presumptive matrix

• not investigated

IRS = inner root sheath; ORS = outer root sheath; HM = hair matrix; SG = sebaceous gland

intercellular adhesion molecules (ICAM), ICAM-1 (CD54), -2 and -3, vascular cell adhesion molecule-1 (VCAM-1), CD2, CD4, CD8, lymphocyte function-associated antigen-3 (LFA-3), the sialoadhesins, and the neural cell-adhesion molecules (NCAM).51-54 NCAM is particularly interesting for hair biology (see below). It is a widely distributed and abundant CAM, which primarily mediates homophilic binding, but heterophilic properties have also been described.55 NCAM is encoded by a single gene, but is expressed as several alternatively spliced forms.56-59 The most common forms, NCAM-120, NCAM-140, and NCAM-180, differ in the size of their cytoplasmic domain and their association with the cell membrane.60,61 NCAM-dependent adhesion is modulated by changes in the amount62 or structure of NCAM,63,64 and can be altered by adding carbohydrate moieties such as polysialic acid (PSA) to NCAM polypeptides.62,65,66 PSA may function as a “cap” on NCAM which inhibits the homophilic binding of NCAM to NCAM on opposed cells. NCAM is

Topobiology of the Hair Follicle

287

expressed in specific spatiotemporal patterns in many tissues during embryogenesis,67-70 and is a key determinant of neuron-Schwann cell and neuron-extracellular matrix interactions during neuronal growth, development, sprouting, remodeling, and regeneration.71-74 NCAM expression is modulated, e.g., by nerve growth factor75 and by several members of the TGF-β superfamily.76-80 Its expression may be controlled by Hox and Pax genes, which implies that it is critically involved in place-dependent pattern formation.81-85 Integrins Integrins are a family of heterodimeric membrane glycoproteins which are composed of an alpha and a beta chain. Fifteen different alpha and eight beta-chains have been reported so far. Originally these adhesion molecules were given the name “integrins” since they integrate the intracellular and the extracellular scaffolds of cells, i. e. cytoskeleton and extracellular matrix. Recently, however, they have been shown to be critically involved in intracellular signaling and the modulation of gene expression.20,85 Cellular binding via integrins is crucial for adhesion and migration on extracellular matrix compounds, and many integrins are components of focal adhesion plates and hemidesmosomes.19,20 Several integrins share on their extracellular side a binding sequence which recognizes the tripeptide arginine-glycine-aspartate (RGD) of other adhesive and matrix proteins,87 and the application of peptides that contain specific RGD motifs can disrupt integrin-mediated binding.88-90 In general, an integrin heterodimer binds more than one ligand, while multiple integrins share ligands and are expressed on the same cell type. Although there is considerable redundancy in the binding capacities of most integrins, most of the integrin mutations reported so far display distinct phenotypes91 (cf. Table 15.2). Integrins are crucial for cell migration, since the cytoplasmic domain of the integrin beta-chain binds to alpha-actinin and talin, which, in turn bind to actin filaments; the contraction of these actin filaments can be used by the cell to move across extracellular matrices.20,92,93 Furthermore, integrins serve a multitude of intracellular signaling functions36,86,94-100 such as an increase in intracellular pH,101-103 in intracellular calcium,104-106 inositol lipid synthesis,107 and the activation of cyclin A.108 In the context of hair biology, it is particularly interesting that several integrins are recognized to protect at least partially against apoptosis.33,34,109,110 For example, c-Myc-induced apoptosis of chicken embryo fibroblasts is suppressed by beta-1 integrins;111 also, soluble anti-beta1-blocking antibodies enhance apoptosis in melanocytes attached to fibronectin, while immobilized anti-beta1 antibodies suppress melanocyte apoptosis.112 In fact, different adhesion receptors may be able to convey distinct apoptosis signals (inhibition or stimulation), depending on cell shape, and costimulation by other signals from the local adhesion milieu.12,13 Given the significance of follicle KC apoptosis in HF morphogenesis and cycling,113-117 it is important to examine selected apoptosis-suppressing integrins as intriguing targets for interfering with HF morphogenesis or apoptosis-driven HF regression (catagen). In this review, we will focus on several integrin subunits and heterodimers for which relevant hair research data are available. General features of these relevant integrin subunits are listed in Table 15.2, while Table 15.3 summarizes their expression patterns during HF development and their potential functional significance for HF morphogenesis.

The “Morphoregulator Hypothesis” A large array of competing and complementary models is now available to explain pattern formation and embryonic periodicity9,11 (see chapter 18)—key themes in HF morphogenesis, with its highly regular and symmetric arrangement of interconnected epithelial cell cylinders. Perhaps the most intriguing among the models explaining the complex role of adhesion molecules in development is the concept proposed by Edelman7,8,84 According

Ligands

• laminin • collagen • fibronectin • tenascin-C

• ICAM-1 • ICAM-2 • ICAM-3 • fibronectin • laminin • collagen • elastase • iC3b • factor X • fibrinogen • kininogen

• laminin

• fibronectin

Integrin Subunit

beta-1

beta-2

beta-4

beta-5

• alternating mesenchymal or epithelial beta-5 expression during tooth morphogenesis205,206 ⇒ Parallels between tooth and HF morphogenesis (cf. chapter 9) encourages investigation of the role of beta-5 integrins in the HF.

• expression in developing tooth epithelium during tooth morphogenesis195 • null mutations display skin blistering disease & perinatal lethality204 ⇒ Parallels between tooth and HF morphogenesis (cf. chapter 9) encourages investigation of the role of beta-4 integrins in the HF.

• beta-2 subfamily of integrins consists of Leukocyte Function Associated Antigen-1 (LFA-1), Mac-1, p150,95198 and alpha-d/beta-2.199 • expressed on most types of white blood cells and crucial elements in inflammatory processes.200-203 • homozygous beta-2-deficient mice show a chronic hyperproliferative inflammatory skin disease approximately 11 weeks after birth, characterized macroscopically by erythema, hair loss, and the development of scales and crusts, which corresponds microscopically to epidermal hyperplasia of the epidermis, subcorneal microabscesses, orthohyperkeratosis, parakeratosis, and lymphocyte exocytosis.180 • beta-2 integrins may be important for the developmental patterning of the HF immune system ⇒ Though the hair loss in beta-2 knock out mice may be secondary to inflammatory skin changes, it deserves to be examined whether beta-2 loss also affects the proliferation or differentiation of HF keratinocytes and the assembly of the HF immune system119

• subunit is shared by all integrins of the fibronectin receptor- or very late antigen-(VLA) family.193,194 • beta-1 expression in tooth epithelium during tooth-development195 • null mutations show peri-implantation lethality183,196 • transgenic mice that express beta-1 integrins in the suprabasal layers of the epidermis develop a psoriasis-like phenotype and hair growth abnormalities.150 ⇒ Parallels between tooth and HF morphogenesis (cf. chapter 9) and psoriasis197 encourages one to investigate the role of beta-1 integrins in HF biology.

• Functional Comments

Table 15.2. Examples of integrins with potential relevance to hair follicle topobiology

288 Molecular Basis of Epithelial Appendage Morphogenesis

• fibronectin

• laminin • collagen

• fibronectin

• laminin

• laminin • tenascin-C

• fibronectin

beta-6

alpha-2

alpha-5

alpha-6

alpha-9

alpha-V

• alternating mesenchymal or epithelial alpha-V expression during tooth development195 ⇒ Parallels between tooth and HF morphogenesis (cf. chapter by 9)

• developmental expression of this integrin coincides with processes of differentiation (alpha-SM actin expression in smooth muscle cells, stratification in epithelial sheets).148 ⇒ alpha-9 shows overlapping expression patterns with NCAM and may play a crucial role in the segregation of DP and CTS fibroblasts.

• alpha6/beta4 is concentrated in hemidesmosomes and has been implicated in the stable attachment of the basal epidermal cells to the basement membrane209,210 • the spatiotemporal distribution and the application of function-blocking antibodies have suggested an essential role in endoderm migration and kidney tubule formation.211-213 • knock out mice show neonatal lethality and a skin phenotype reminiscent to epidermolysis bullosa.214 ⇒ The epidermolysis bullosa-like skin phenotype suggests that this integrin subunit is essential for HF attachment to its basement membrane.

• knock out mice display pronounced defects in posterior trunk and yolk sac mesodermal structures and embryonic lethality.208 • transgene mice with suprabasal epidermal overexpression (α5, α5β1) displayed a psoriasis-like phenotype150 ⇒ Parallels between HF morphogenesis and psoriasis197 encourages investigation of the role of alpha-5 integrins in HF biology

the HF.

• transgenic mice with suprabasal epidermal α2β1-overexpression displayed a psoriasis-like phenotype150 ⇒ Parallels between HF morphogenesis and psoriasis197 encourages investigation of the role of alpha-2 integrins in

• highly expressed in inflammatory processes and in malignant epithelial neoplasms.207 • knockouts display cutaneous macrophage infiltration and juvenile baldness149 ⇒ Though the hair loss in beta-6 knock out mice may be secondary to the macrophage infiltration and inflammatory cytokines, beta-6 loss might also affect the proliferation /differentiation of HF keratinocytes and the assembly of the HF immune system119

Topobiology of the Hair Follicle 289

transgenic mice with suprabasal overexpression (α2β1) had fewer whiskers and a psoriasis-like phenotype150

accumulation of immune cells close to the hair canal transgenic mice with suprabasal overexpression had fewer whiskers (β1 alone,α2β1 or a5β1), disturbed orientation of back skin hairs (β1) and the whiskers were short and curly (β1); psoriasis-like phenotype (β1, α2β1, α5β1)150

ORS in human HF215

ORS of human HF; similar staining patterns in fetal, newborn, adult human skin215

IRS, yet only in fetal skin215

developing hair placode and hair bud KC. Intense IR on the future DP and CTS cells148

LC/MAC in the distal ORS during neonatal murine HF development

ORS (adolescent human skin)215

alpha-2

alpha-3

alpha-5

alpha-9

alpha-E

beta-1

IRS = inner root sheath; ORS = outer root sheath; HF = hair follicle; HM = hair matrix; LC = Langerhans cells; SG = sebaceous gland

overlapping expression pattern with NCAM

transgenic mice with suprabasal overexpression (α5, α5β1) had fewer, shorter and curly whiskers and disturbed orientation of back skin hairs and a psoriasis-like phenotype (α5β1)150

Comments

Expression Pattern Subunit

Integrin

Table 15.3. Expression patterns of selected integrins potentially relevant to hair follicle development

290 Molecular Basis of Epithelial Appendage Morphogenesis

Topobiology of the Hair Follicle

291

to his so-called “Morphoregulator (MR) Hypothesis”, one essential link between regulatory genes and three-dimensional pattern formation is the coordinated expression and function of adhesion molecules, referred to as MR molecules7,8,84 (Fig. 15.1). Edelman has proposed their expression to be controlled by three kinds of genes: by the genes coding for the above MR molecules (MR genes); by so-called “historegulatory” genes (HR genes), i. e., genes which regulate all other cytodifferentiation events not related to cell shape, pattern formation and migration; and by so-called “selector” genes such as homeobox (Hox) and paired box (Pax) genes, which restrict the expression of both HR and MR genes to certain developmental places and times. According to this hypothesis, inductive signals lead to the expression of particular combinations of MR molecules which mediate the establishment of cell borders in a collective of functionally linked cells.7,8,84 Following the “MR Hypothesis”, regulatory loops get initiated when a morphogen secreted by one cell collective induces another cell collective to enter on a specific developmental path associated with a distinct pattern of adhesion molecule expression, leading to differences in cell shape, polarity, migration, proliferation, and/or apoptosis. The second cell collective may now release a second or multiple morphogens that, in turn, send additional inductive signals to the first cell collective, causing another change in the local adhesion milieu, and so forth84 (Fig. 15.1). Since this may well be the most comprehensive and convincing concept on the role of adhesion molecules in morphogenesis proposed so far, it offers a useful theoretical framework for exploring the functions of adhesion molecules in HF development. In the following, we shall therefore adhere to this hypothesis when interpreting the scarce data that are currently available on HF topobiology (Table 15.1, 15.3).

Topobiological Questions in Hair Follicle Morphogenesis As outlined in chapter 6, the morphogenesis of HFs—one of the defining features of mammals—requires the invagination and specialization of rather undifferentiated epithelial cell populations (epidermal KC) within a specialized mesenchymal environment (i.e., fibroblasts of the future perifollicular connective tissue sheath and the dermal papilla) (cf. Figs. 15.2-15.4 and chapter 6. This epithelial invagination develops in order to form a multicylindric, highly differentiated, keratinized epithelial reduplication (hair shaft and follicle sheaths) which protrudes far beyond, and imparts substantial additional functions to, the epithelium from which it has originated3,117,118 (chapter 7). Topobiologically, this begs many pertinent questions, such as: Which MR molecules are critically involved in hair placode formation and define the boundaries of the hair placode even before it becomes recognizable by morphological criteria?1,119 How do MR molecules instruct and coordinate the downward migration of KC within the developing epithelial hair bulb during the first three stages of HF morphogenesis? How are specific MR molecules exploited to instruct the majority of proximal epithelial hair bulb KC to change their migratory pattern from a distal-to-proximal to a proximalto-distal direction (with the formation of the inner root sheath and the hair matrix, the previously “epidermofugal” growth direction of developing hair bulb KC just above the future dermal papilla suddenly becomes “epidermopetal”) (cf. chapter 6)? How are specific MR molecules exploited to dictate KC segregation into distinct subpopulations that subsequently form the HF sheaths (e.g., outer and inner root sheath, cuticle) all of which differ substantially in their characteristics of proliferation, metabolic and secretory activities, terminal differentiation, and in the pattern of programmed cell death3,116,119-121 (chapter 7)? Which MR molecules are critically involved in segregating melanocyte and sebocyte precursors, hematopoietic cells, epithelial stem cells, transient amplifying cells, and terminally

Molecular Basis of Epithelial Appendage Morphogenesis

292

Binding Cell Collective B Selector Genes

Induction

Induction

Selector Genes

Binding Cell Collective B

Fig. 15.1. The “Morphoregulator Hypothesis”. According to this hypothesis, the coordinated expression and function of adhesion molecules, referred to as MR molecules, is controlled by three sets of genes: by the genes coding for the above MR molecules (MR genes); by “historegulatory” genes (HR genes) that regulate cytodifferentiation, and by “selector “ genes such as homeobox (Hox) genes which coordinate the precise spatiotemporal distribution of HR and MR gene products. Regulatory loops get initiated when a morphogen secreted by one cell collective induces another cell collective to enter on a specific developmental path associated with a distinct pattern of adhesion molecule expression, leading to differences in cell shape, polarity, migration, proliferation, and/or apoptosis. The second cell collective may now release a second or multiple morphogens that, in turn, send additional inductive signals to the first cell collective, causing another change in the local adhesion milieu, and so forth. (Modified after Edelman8,84).

Topobiology of the Hair Follicle

Fig. 15.2. E-cadherin expression during neonatal murine hair follicle morphogenesis. Strong E-cadherin IR was found on keratinocytes of the epidermis and the developing hair placode, the outer root sheath (ORS), the inner root sheath (IRS), the outermost hair matrix (OHM), and the myoepithelial cells which surround the sebocytes of the sebaccous gland (SG); weak E-cadherin IR was noted in the keratogenic region of the hair matrix. The bulge region, the putative stem cell compartment of the hair follicle, displayed heterogenous E-cadherin IR. d = dermis, sc = subcutis, DP = dermal papilla. Stages of HF morphogenesis adopted after Hardy.1 Details: see chapter 6. (Modified from ref. 124).

293

294

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 15.3 P-cadherin expression during neonatal murine hair follicle morphogenesis. Intermediate P-cadherin IR has been found on keratinocytes of the epidermis and of the developing hair placode. Strong P-cadherin IR was noted on the keratinocytes of the developing outer root sheath (ORS), the innermost hair matrix (IHM), and the keratogenic region of the hair matrix. d = dermis, sc = subcutis, DP = dermal papilla. Stages of HF morphogenesis adopted after Hardy.1 Details: see chapter 6. (Modified from ref. 124).

Topobiology of the Hair Follicle

295

Fig. 15.4. NCAM expression during fetal murine hair follicle morphogenesis. Strong epithelial NCAM IR has been detected on hair placode keratinocytes (KC) during stage 2; it became restricted to KC of the future outer root sheath (ORS) during stage 3 and to peri- and suprainfundibular KC during stage 5 to 8. Weak mesenchymal NCAM IR has been noted on all dermal cells before placode formation (not shown), while strong ubiquitous dermal NCAM IR was found during stages 4 to 8 (gray underlaid box). (Modified from ref. 135).

differentiating KC from each other within the developing follicle epithelium, and how are MR molecules employed to determine the final location of these cells within the mature, growing HF? To which extent are MR molecules responsible for the developmentally crucial condensation and morphogen secretion of those dermal fibroblasts that specialize to become the follicular papilla (DP) and the perifollicular connective tissue sheath (CTS)? Which MR molecules provide the developmental basis for the astonishing symmetry and periodicity of the defined epithelial cell compartments that characterize the mature, growing HF in its anagen VI state? How can the targeted manipulation of homophilic and heterophilic interactions of MR molecules be therapeutically exploited to assist in the induction of de novo HF morphogenesis and/or in modulating the cycling, or even the ablation, of HFs in the adult mammalian organism? None of these questions, which define the major challenges carved out for “trichotopobiology” can presently be answered satisfactorily. However, the advent of powerful new molecular research tools, namely the ever-increasing number of mouse mutants with knockout or targeted overexpression of defined adhesion molecules, and the availability of well-defined hair research models, especially of murine models that offer valuable background data for the interpretation of the HF observations made in adhesion mutants, promise that some of these questions may soon be answered.

296

Molecular Basis of Epithelial Appendage Morphogenesis

Trichotopobiology In addition, for a few selected MR molecules suggestive evidence has been accumulating that they are indeed important elements in the control of HF morphogenesis (see below). Yet (as already hinted in Tables 15.1-15.3), our knowledge of trichotopobiology is mostly restricted to some phenomenological observations, and we are still very far from being able to synthesize a comprehensive and convincing picture of exactly how specific MR molecules bring about their developmental effect in HF biology. In fact, hair researchers have not yet acquired the level of topobiological insight that their colleagues in feather and tooth morphogenesis have won over the last years (cf. chapters 5, 9). Nevertheless, at least with respect to E- and P-cadherin, NCAM, and selected integrins, some basic data are now available and a first working hypotheses can be formulated. E- and P-cadherin Ecad and Pcad expression has been studied in several skin and HF types during murine49,122-124 (Figs. 15.2, 15.3) as well as human50,125 morphogenesis (summarized in Table 15.1). Notably, in the skin both cadherins are detected only on epithelial, but not on dermal cells, which is one of the reasons why these cadherins are considered critical determinants of the “epithelial” phenotype.27-32,126 Generally speaking, Ecad is initially present on all epithelial cells, and Ecad has been found in all layers of murine and human epidermis. During the consecutive stages of murine HF development, Ecad is progressively downregulated on the developing inner hair matrix (IHM) and inner root sheath (IRS) (summarized in Fig. 15.2).49,50,122-125 Strong Ecad expression is seen in the suprabasal epidermal layers, the ORS, the IRS, and the outer part of the hair matrix (OHM).49,50,122-125 During human and murine fetal HF development,50,122 Pcad was found to be expressed by the KC of the basal epidermal layer and of the developing IHM, IRS, and ORS. The latter is continuous with the basal epidermal layer. Ecad is also expressed by the myoepithelial cells of the murine sebaceous gland. Interestingly, in neonatal murine and fetal human skin, Ecad expression is weaker in the Pcad+ basal layer compared to the suprabasal epidermal layer.50,123,124 In contrast, in the basal layer of hairless human skin of the palm and foreskin, patches of brightly labeled cells have been detected with antibodies to Ecad, beta-catenin, and gamma catenin, but not with antibodies to Pcad, alpha-catenin, pan-desmocollin and pan-desmoglein.48 The authors suggest that these inhomogenous expression patterns may indicate gradients of distinct cell-cell and cell-matrix adhesiveness which may provide markers for the stem cell compartment.48 Further studies are needed to elucidate whether similar gradients can be found in the bulge area, the putative stem cell compartment of the HF.127-130 During the neonatal development of murine pelage follicles (cf. chapter 6), we noted intermediate levels of Pcad IR on all epidermal and hair placode KC. Pcad IR became progressively restricted to the ORS and the innermost portion of the developing hair matrix (OHM), at the same time that Ecad expression became restricted to the KC of the epidermis, the ORS and the outermost portion of the hair matrix (OHM). Briefly, Ecad IR was not detected on the KC of the IHM, while Pcad IR was not noted on the OHM and IRS and was found more weakly on KC of the epidermis and the suprainfundibular region of the HF compared to ORS and IHM (Figs. 15.2, 15.3).123,124 That study revealed at least four distinct patterns of follicular Ecad/Pcad expression during murine HF morphogenesis: 1. Isolated P-cad expression—inner layers of the hair matrix; 2. Isolated Ecad expression—inner root sheath; 3. Combined strong expression of Ecad and Pcad—outer root sheath; 4. Strong Pcad expression combined with weak Ecad expression—keratogenic region of the precortical hair matrix

Topobiology of the Hair Follicle

297

It is tempting to speculate that these distinct topobiological patterns reflect follicular KC segregation by Ecad and Pcad into defined subpopulations of HF cells. For example, isolated Pcad expression (pattern 1) may support the sorting out of that subpopulation of HF KC which are in direct contact with the DP and which may be (or might become) susceptible to morphogenic signals from the DP during hair growth induction. Strong Pcad expression combined with weak Ecad expression (pattern 4) may, instead, contribute to the segregation of precortical HM KC which are programmed to enter into terminal differentiation and trichocyte formation in the keratogenic region of the anagen HF. Pcad colocalizes with the proliferation marker Ki-67 in the epidermis and in IHM KC;50 consequently, it has been proposed that Pcad may be essential to segregate the proliferating basal epidermal layer from the differentiating upper layers, although Ecad is also needed for the fundamental linking of these layers.122 Thus, Pcad may serve to segregate proliferating KC of HFs and epidermis from the differentiating cell pool. Cadherin-mediated signaling has been implied in the control of terminal differentiation in epithelial cells, since antibodies to Ecad and Pcad, which block calcium-induced stratification, prevent the selective loss of integrins (mRNA and protein) that is associated with KC terminal differentiation.131 The introduction of a dominant negative E-cadherin mutant, which lacks the extracellular domain of Ecad, into normal human KC via a retroviral vector reduces KC proliferation, desmosome formation, and stratification as well as integrin expression.132 This strongly supports the concept that cadherin-mediated signal transduction events are a key element in controlling the terminal differentiation of epithelial cells. With this background, it now needs to be elucidated whether the distinct expression patterns of Ecad and Pcad during HF morphogenesis (Figs. 15.2, 15.3) are of secondary or primary importance: Is combined weak Ecad and strong Pcad expression causally involved in inducing the apparently highly controlled switch from proliferation (proximal hair matrix) to differentiation (keratogenic precortical matrix region, hair shaft cortex)? Interestingly, Hirai et al122 have demonstrated that the administration of function-blocking antibodies against Ecad and Pcad disrupt epithelial tissue architecture in cultured pieces of murine embryonic upper lip skin and interfere with pattern formation of a subpopulation of dermal fibroblasts, which—like all fibroblasts—was cadherin-negative. In contrast to untreated control lip skin pieces, which displayed normal patterns of HF morphogenesis, skin cultures treated with monoclonal antibodies to Ecad (ECCD-1) or to P-cadherin (PCD-1) showed abnormal HF morphogenesis with the following features: HFs were deformed; the cuboidal or columnar arrangement of epidermal basal layer cells was distorted; and the condensation of the perifollicular dermal cells was suppressed, causing an unusually homogeneous distribution of these fibroblasts. The latter phenomenon was more pronounced after PCD-1 than after ECCD-1 treatment, and strongest after simultaneous blocking of Ecad and Pcad.122 A similar effect was found by using antibodies against the chicken homologue of Ecad (L-CAM) in chicken skin explant cultures:133 highly specific antibodies to epithelial L-CAM disturbed dermal pattern formation, causing a nonhexagonal striped pattern of the dermal condensations. Thus, the disruption of normal epithelial linkage phenomena by function-blocking anti-cadherin antibodies in avian and murine skin perturbed pattern formation of selected, constitutively cadherin-negative dermal cells. This suggests that the production of inductive signals from the epithelium to the mesenchyme depends on the integrity of cadherinlinked epithelial cell collectives.122,133 The establishment of Ecad and Pcad-mediated cell junctions may be crucial for the basal epidermal and/or ORS KC to produce appropriate signals for the perifollicular dermal condensation, since the simultaneous disruption of Ecad and Pcad linkage had the biggest perturbing effect on dermal pattern formation.122

298

Molecular Basis of Epithelial Appendage Morphogenesis

Topobiologically, these findings not only suggest that cadherins may control epithelial cell shape (distorted cell shape of basal KC) and the three-dimensional architecture of supracellular structures (deformed HFs), but also raise the possibility that cadherins link KC subpopulations into functional units with “inductive” properties on the associated mesenchyme (disrupted condensation of perifollicular dermal fibroblasts). The mesenchyme, in turn, would then fail to generate the appropriate signaling response in this complex epithelial-mesenchymal interaction system1 (chapter 6), thus producing the morphological abnormalities seen in the epithelial tissue compartment. In summary, Ecad and P-cadherin show spatiotemporally controlled and apparently stringently coordinated expression patterns on distinct subpopulations of epidermal and hair follicle KC during murine and human HF development (Figs. 15.2, 15.3, Table 15.1). Pcad may be essential for segregating the proliferating and differentiating KC pools in the HF, while Ecad may be the “basal glue” linking KC into defined epithelial sheets. We propose that both cadherins are essential for the segregation of follicular KC into the ORS (Ecad+, Pcad+), IRS (Ecad+), and the keratogenic precortical matrix region (weakly Ecad+, strongly Pcad+), and indirectly regulate the induction of the perifollicular dermal fibroblasts condensation. NCAM Among the multitude of members of this very large family of cell-adhesion molecules, data relevant for hair biology are only available for ICAM-1125 and NCAM.4,123,125,134,135 Since the data concerning ICAM-1 expression during HF development are rather limited (transient expression of ICAM-1 on the outer cells of hair germs (stage 2) in fetal human skin at 11-12 weeks of estimated gestational age125), and since we failed to detect ICAM-1 expression on murine HF before the first anagen V of HF morphogenesis in neonatal C57BL/6 mice,119 we will focus here on NCAM. The fundamental functional importance of NCAM for skin appendage formation was first established by Chuong and co-workers, who studied developing feather follicles in chicken skin explant cultures.4-6,136 As reported and discussed in greater detail in chapter 13, the authors demonstrated that NCAM-mediated signaling events are crucial to the appropriate formation of the dermal condensations next to developing follicle buds: the application of anti-NCAM antibodies during feather follicle morphogenesis in vitro induced a highly irregular shape of the dermal condensations, i.e., giant, tiny or missing buds were found, compared to the physiological hexagonal pattern.136 Since the morphogenesis of feather and hair appears to share many developmental controls and follows similar developmental principles (chapters 1, 5, 6, 13, 18-20), similar NCAM expression patterns in feather and HF4 strongly encourage one to investigate the role of NCAM in trichotopobiology, particularly during the development of the perifollicular CTS and the DP. During fetal HF development in mice, NCAM expression has been reported in the DP, the CTS, and the ORS.4,135 Vielkind et al135 have demonstrated that NCAM expression in C57BL/6 mice is spatiotemporally highly controlled during the development of tylotrich HFs in fetal skin. Figure 15.4 summarizes the transient follicular and perifollicular NCAM expression patterns during defined stages of the development of tylotrich and pelage HF as described by Vielkind et al.135 Before the development of the hair placode (stage 1), some NCAM staining was found on all cells of the future dermis. During hair placode development, NCAM IR was noted only on selected fibroblasts of the papillary dermis that was in close vicinity to the epidermis, while NCAM IR disappeared from the remainder of the dermis. During hair peg formation of tylotrich HF, most dermal cells in close vicinity to the epidermis as well as most epithelial cells of the hair pegs were NCAM+. During the DP morphogenesis, epithelial

Topobiology of the Hair Follicle

299

NCAM IR became more and more restricted to the isthmus region of each follicle as well as to a few perifollicular basal layer epidermal KC. Furthermore, dermal NCAM IR was noted again, in particular on the fibroblasts of the CTS and on the fibroblasts of the DP. During the early development of the IRS, the DP, the proximal CTS, and the mesenchymal cells in the distal interfollicular dermis showed increasingly intense NCAM IR.135 Using the same antibody and the same mouse strain,137,138 we have largely reproduced these NCAM IR patterns, but have noted a significant change of NCAM IR pattern during murine HF morphogenesis, when HF development that occurred in late fetal skin was compared to HF morphogenesis in neonatal mouse skin. Compared to late fetal skin and HF development, NCAM IR was even stronger throughout the dermis during neonatal morphogenesis of murine pelage HF(day 1 -21 post partum). Similar to fetal NCAM expression, the entire CTS and the DP fibroblasts during stages 5 to 8 showed strong NCAM IR, as did the KC of the supra- and periinfundibular region of the developing ORS. Later, the initially strong, ubiquitous dermal NCAM IR became progressively restricted to the fibroblasts of the future DP and of the entire perifollicular CTS.137,138 However, the dermal NCAM expression pattern during fetal human HF morphogenesis reportedly is similar to the one seen in neonatal murine HF development. NCAM is initially present in the entire dermis and gets progressively restricted to the dermal condensation, which gradually develops into the DP, and the perifollicular CTS.125 This raises the intriguing possibility that the NCAM expression pattern of embryonic human dermis and HFs reflects the installation of similar developmental programs as those operating during neonatal murine skin development. The strong ubiquitous dermal NCAM expression during human and neonatal murine pelage HF development may be due to a precise spatiotemporal regulation of distinct NCAM isoforms, which cannot be distinguished by the employed antibody. The strong dermal NCAM expression may reflect the presence of an NCAM-upregulating, local cytokine or neurotrophin milieu (e.g., NGF or members of the TGF family modulate NCAM expression).75-80 Such a local signaling millieu favoring mesenchymal NCAM expression may only exist during HF and skin morphogenesis and may be evolutionarily conserved between men and mice. Topobiologically, the spatiotemporal distribution of NCAM expression during HF morphogenesis, as well as the functional effects of NCAM-blocking antibodies on feather development, raise the question of whether NCAM may be utilized as a key downstream effector molecule for the stringently controlled bidirectional epithelial-mesenchymal inductive processes during HF development1 (chapter 6). DP and CTS fibroblasts have the unique capacity to induce hair growth.139-141 As Jahoda and Reynolds141 have pointed out, not only the DP fibroblasts themselves, but also their structural relationships to one another, to hair matrix KC, and to CTS fibroblasts are important elements in HF growth control. Namely, the germinative cells of the HF epithelium are surrounded by and interact with two distinct dermal cell populations, i. e. fibroblasts of the DP and the proximal CTS; these special fibroblast populations may cooperate to elaborate a two-sided gradient of mesenchymal morphogens which controls germinative cell activities.141 Although this hypothetical model has been developed for adolescent anagen HF, it may be instructive to apply it also to HF morphogenesis, since adolescent HF cycling, specifically the telogen-anagen transformation, has long been thought to recapitulate many aspects of HF morphogenesis.1,3,118,120,121 If one assumes that the condensation of the later CTS and DP fibroblasts early during HF morphogenesis is important to segregate and differentiate dermal fibroblasts into morphogen-elaborating cell collectives which thereby attain novel functional properties compared to neighboring fibroblasts, it is tempting to speculate that NCAM is a key adhesion molecule for this fibroblast segregation and differentiation and the subsequent

300

Molecular Basis of Epithelial Appendage Morphogenesis

generation of a two-sided, mesenchyme-derived morphogen gradient. This speculation is prompted by the striking congruence between the distribution of putatively morphogen gradient-generating CTS and DP fibroblasts with the NCAM expression pattern during HF morphogenesis: as Figure 15.4 illustrates, in and around the proximal HF, NCAM expression becomes progressively restricted to the CTS and DP as HF development advances. Since NCAM is a crucial factor for axonal outgrowth by activating the neuronal fibroblast growth factor receptors via phosphorylation,142 the purpose of NCAM expression on fibroblasts may go well beyond that of mechanical cell sorting functions. For example, it is conceivable that the NCAM expression on DP and CTS fibroblasts also induces a higher susceptibility of the mesenchymal cells to morphogens and/or inductive signals released by the proximal HF epithelium. Integrins A large number of recent mutations of most integrin genes91 has recently provided novel insights into the key role of defined integrins in many developmental systems, such as the morphogenesis of kidneys,143 placenta and heart.144 In contrast, the functional significance of most integrins in HF morphogenesis remains obscure, even though an increasing number of integrins is reported to be expressed in the HF. Table 15.3 summarizes the expression patterns during HF morphogenesis that have been reported so far, as well as published HF phenotypes of integrin null mutations or targeted overexpression. Conclusive functional analyses have been hampered by the fact that the mutations of several integrins which are predominantly expressed in the mesenchyme are associated with embryonic lethality,91 i.e., preclude the analysis of HF which could have developed only thereafter. Some epithelial integrins which bind to basement membrane components display milder phenotypes than the mesenchyme-associated integrins, which points to the existence of partially redundant, overlapping functions of several integrins and the operation of successful compensatory mechanisms in several integrin null mutations.91 Unfortunately, none of these integrin mutants with milder phenotypes have been rigorously and systematically screened, employing established hair research techniques, for abnormalities in HF morphogenesis compared to normal HF development (see chapter 6), specifically in the respective wildtype littermates.145-147 The most detailed integrin expression data are currently available for the alpha-9 integrin subunit, which only forms an integrin heterodimer with the beta-1 subunit. The only ligand reported so far is tenascin-C. Expression of alpha-9 integrin and of its ligand, tenascin-C, has been noted to coincide with stratification and cornification of embryonic murine epidermis. The developing hair placode shows strong alpha-9 IR, which remains restricted to hair bud KC during the subsequent follicle elongation.148 The developing ORS displays no alpha-9 IR, while tenascin-C IR is detectable during the onset of follicular growth. Intense alpha-9 IR is found on the future DP and CTS cells. In contrast to NCAM IR, the most pronounced tenascin-C IR is seen on distal CTS fibroblasts close to the epidermis, and reportedly absent in the mature DP.148 Since the expression of alpha-9 occurred in conjunction with the development of the differentiated phenotype,148 it has been proposed that this expression pattern of alpha-9/beta-1 correlates with the maintenance of the mature phenotype of organs rather than with early steps in tissue morphogenesis.148 Beta-6 deficient mice display cutaneous macrophage infiltration and show at day 5 postpartum macroscopically reduced hair development (heads, neck region, and ventral thigh skin), yet only in selected regions of the integument.149 The mice had fewer HF, and numerous degenerating HF surrounded by mononuclear cells. The involucrin promoter-driven overexpression of several integrin subunits (alpha-2, alpha-5, beta-1) in suprabasal layers of the epidermis and the ORS induces a psoriasis-like

Topobiology of the Hair Follicle

301

skin phenotype, as well as whisker and pelage HF abnormalities including a reduced total number of whisker follicles as a particularly frequent observation.150 Mice overexpressing alpha-5, beta-1 or both integrins display a disturbed orientation of HFs in all regions of the integument which have been examined, as well as shorter and curly whiskers. Interestingly, the photodocumentation of back skin shows telogen follicles in wildtype skin sections as opposed to anagen follicles in skin sections of age-matched transgenic animals.150 A professional and systematic trichological analysis of these transgenic mice, therefore, promises to clarify the influence of specific integrins in the suprabasal KC of epidermis and follicle ORS on murine HF development and cycling.

Control of Adhesion Molecule Expression During Hair Follicle Morphogenesis In search of the “selector” genes According to the MR hypothesis,7,8,84 homeotic genes, which have a conserved motif in the homeobox (Hox) in common and code for proteins with transactivating gene regulatory functions,151,152 are good candidates for the role of chief controllers of MR molecule expression during skin appendage formation6,70,85 (see chapters 4, 5). Hox genes control the developmental fate of cells by encoding homeoproteins that bind specific DNA sequences and regulate gene expression at the level of transcription.151,152 The term “homeodomain” defines a class of protein domains that bind to a defined group of promoters and have a recognizable similarity to a 60 amino acid motif which is encoded by a 180 bp homeobox sequence.151-153 Homeotic genes play a pivotal role in controlling the three-dimensional pattern formation of tissues and organs of invertebrates and vertebrates, and specify the attributes of entire body regions. They are positioned on the chromosome in a sequence that corresponds to the spatiotemporal expression pattern of these homeotic genes during development of the mammalian body, and the emission as well as the response to inductive signals result in changes of the Hox gene expression.11,19,151,152 Not surprisingly, many Hox gene mutations cause incorrect specification of body parts11,151,152,154-156 (see chapter 18). In general, single mutations in Hox genes appear to lead only to fairly subtle abnormalities; however, combined mutations of more than one Hox gene lead to gross abnormalities in multiple organ systems157 (M. R. Capecchi, personal communication), including skin and HFs (M. R. Capecchi, personal communication). According to the MR hypothesis,7,8,84 specific “selector” genes sequentially turn on or off effector genes (in this context referred to as HR and MR genes) in defined cell collectives (Fig. 15.1). Good examples are several Hox and Pax gene products which bind and regulate the NCAM promotor in vitro, e.g., the products of HoxB8, HoxB9, Hox C6, Phox-2, cux, Pax8.70,81-83 These “selector” genes may thus control the spatiotemporal distribution of these MR molecules to segregate follicular KC or dermal fibroblasts into functionally distinct subpopulations (e.g., ORS, IRS or DP, CTS, respectively) as well as to guide the acquisition of new functional properties such as the elaboration of morphogens. Hox genes are organized in a hierarchical system and show macrogradients as well as microgradients5,151,152 (see chapter 5). A macrogradient represents a position-specific expression of homeoproteins along the body axis, e.g., a strong expression of a homeoprotein in the tail region which is absent in the neck region.158 A microgradient has been reported in feather buds,159 i.e., similarly to the macrogradients a strong homeoprotein expression can be detected in the cephalic part which is absent in the caudal part of a feather follicle.159 Such a homeoprotein gradient might be a key element in establishing the anterior-posterior axis of skin appendages.5,6

302

Molecular Basis of Epithelial Appendage Morphogenesis

Only a few Hox genes have been studied so far in mammalian skin and its appendages.5,6,160-169 Since “selector” genes themselves are usually tightly embedded into a network of positive and negative controls,11,153 in the context of trichotopobiology, it is of particular interest to define specific regulatory loops between MR and selector genes and the proteins coded by them. One such regulatory loop, which might be important for trichotopobiology, is the interaction between NCAM and putative “selector” genes. As mentioned before, several Hox and Pax gene products bind and modulate the NCAM promotor81-85 and may regulate the spatiotemporal distribution of NCAM during development. Furthermore, NCAM activates FGF receptors,142 which leads to a higher susceptibility of neuronal cells to stimulation by members of the FGF family. The members of the FGF family have, in turn, multiple influences on several “selector” genes.170-173 Another loop can be envisioned for interactions between Ecad and lymphocyte enhancer-binding factor-1 (Lef1). The transcription factor Lef1 plays a fundamental role in the formation of HFs and several other organs and tissues that depend on inductive epithelial-mesenchymal tissue interactions174,175 (chapter 6). This is illustrated by the fact that Lef1-deficient mice lack teeth, mammary glands, whiskers, and a fur coat.176 Lef1 and betacatenin form a ternary complex with DNA that displays an altered DNA bend and regulates gene expression by direct binding.46 Beta-catenin, in turn, links cadherins to the cytoskeleton, while the beta-catenin/Lef1 complex associates with the 5' end of the Ecad gene, whose expression it may modulate,47 thus regulating Ecad expression. Thus, the interaction of Lef1 and beta-catenin may provide a trichobiologically pivotal molecular signal transduction loop, from cell-adhesion at the cell surface to nuclear gene expression, and back to the expression of adhesion molecules.

Perspectives On the basis of these limited data and considerations, it is possible to develop a hypothetical scenario that attempts to explain how Ecad, Pcad, and NCAM may be exploited for the generation of pattern formation during murine HF morphogenesis (Fig. 15.5). An as yet unknown “first message” of ill-defined origin is widely thought to instruct a defined sector of the cutaneous epithelium to make a skin appendage1 (chapter 6). According to the MR hypothesis,7,8,84 these morphogens turn on or off epithelial selector genes. These, in turn, control the defined spatiotemporal expression of genes responsible for the proliferation or differentiation of selected KC (HR genes) in specific KC subpopulations, or for the coordinate expression patterns of MR molecules such as Ecad or NCAM (MR genes). The binding of the newly expressed MR molecules to their ligands, such as homophilic binding of Ecad to Ecad, may lead to a segregation of the proliferating hair placode KC from epidermal KC. In addition, this may induce intracellular signal transduction cascades (such as the putative cascade Ecad → beta-catenin → Lef1 → Ecad) which alters directly or indirectly the epithelial gene expression and upregulates the elaboration of inductive molecules such as members of the TGF-β/BMP and/or FGF families of growth factors177 (chapter 6). According to this hypothetical scenario (Fig. 15.5), these inductive molecules may subsequently turn on or off dermal selector genes which, in turn, control the defined spatiotemporal expression of HR and MR gene products in specific fibroblast subpopulations, thus controlling e. g. the NCAM expression on selected dermal fibroblasts. The binding of MR molecules to their ligands (such as the homophilic binding of NCAM to NCAM) may then lead to a condensation of selected fibroblasts—the future DP and CTS. Furthermore, this binding of MR molecules to their ligands could subsequently alter the gene expression of DP and/or CTS fibroblasts, which may change the signals emitted by the developing HF epithelium. The NCAM-dependent alterations in mesenchymal gene expression may then trigger the secretion of morphogens from the DP and CTS, which, in turn, regulate the

Topobiology of the Hair Follicle

303

Principles

Details (A) mesenchymal (?) secretaion of inductive molecules (e.g., BMP-2, BMP-4, FGF-4)

(B)

Segregation of proliferating hair placode keratinocytes from epidermal keratinocytes

(B) secretion of inductive molecules

(C) condensation of selected dermal fibroblasts (DP & CTS) & secretion of inductive molecules

(D) segregation of distinct epithelial subpopulations via selective expression of

• strong E- & P-cadherin expression in the ORS • isolated E-cadherin in the outermost hair matrix and IRS • isolated P-cadherin expression in the innermost hair matrix • strong P-cadherin and weak E-cadherin expression in the keratogenic region of the hair matrix

Fig. 15.5 Topobiology of the hair follicle. Simplified working hypothesis on the role of selected adhesion molecules in pattern formation during murine hair follicle morphogenesis. The left column summarizes basic principles of hair follicle morphogenesis according to Hardy1 (A-C) and shows the expression patterns of selected morphoregulatory molecules (NCAM, E- and P-cadherin) during hair follicle morphogenesis (A-D). The right column specifies these basic principles in more detail in a corresponding scenario of topobiological events following the “Morphoregulator Hypothesis”.8,84 Gray background boxes in the right column correspond to selected stages of hair follicle development shown in the left column. See text for details. Abbreviations: ↓, induces/stimulates; CTS, connective tissue sheath; DP, dermal papilla; IRS, inner root sheath; KC, keratinocytes; NCAM, neural cell-adhesion molecule; ORS, outer root sheat.

304

Molecular Basis of Epithelial Appendage Morphogenesis

expression of epithelial selector genes that, finally, control epithelial HR and MR genes. Following this hypothesis, the proliferating KC are now segregated into several subpopulations via selective expression of MR molecules (e.g., combined strong Ecad and Pcad expression in the ORS, isolated Ecad expression in the outermost part of the hair matrix, isolated Pcad expression in the innermost hair matrix, and weak Ecad and strong Pcad expression in the precortical keratogenic region of the hair matrix.) This preliminary working hypothesis offers a useful theoretical framework for the design of the kind of experiments badly needed in order to bring trichotopobiology at least to the level of molecular understanding already achieved concerning the MR controls of tooth and feather (chapters 1, 5, 9, 13, 18-20). A few examples of long overdue key experiments shall suffice in order to elucidate the functional role of MR molecules in HF morphogenesis. One basic and critically important challenge is the systematic analysis of morphological aberrations and the acceleration/retardation of fetal and neonatal HF development in all potentially relevant mouse mutants of adhesion molecules or their ligands. For example, viable null mutants are now available for NCAM,178,179 beta-2 integrins,180-181 and beta-6 integrins149 (see also Table 15.2). Chimeric mice with a mosaic pattern of beta-1-deficient cells are also available.183 Specifically, we advocate performing standardized histomorphometric analyses of differences between wildtype and mutant mice with respect to: 1. HF morphogenesis; 2. spontaneous HF cycling; and 3. experimentally manipulated HF growth and cycling in vivo and in skin organ culture.119,146,147,184-188 It deserves to be emphasized that such analyses are well-advised, irrespective of whether or not the laboratory from which the mutant in question has originated has reported any hair growth abnormalities. Professional hair growth analyses are routinely ignored or neglected in most mutant strains, and—after careful analysis—surprising, ever-increasing numbers of mouse mutations indeed turn out to display (often only subtle) hair growth abnormalities.145 Therefore, we subscribe to the simplistic and provocative credo, “Hair growth in a mouse mutant is abnormal until rigorously proven otherwise”. The targeted “forced” overexpression of epithelial cadherins in defined epithelial compartments35 such as the basal epidermis and the ORS e.g., via the K14 promoter,189 or in the suprabasal layers e.g., via the involucrin promoter,150 will be another key achievement for probing the putatively causal role of Ecad and Pcad in regulating KC shape, segregation, proliferation, and apoptosis and differentiated functions in the HF. As mentioned above, transgenic mice which overexpress integrins are now available, show an impressive skin phenotype reminiscent of psoriasis,150 and could easily be subjected to the analyses suggested above. Other additional pragmatic, but instructive functional experiments can be envisioned in our context. In order to elucidate whether NCAM expression on follicular fibroblasts is a secondary effect or causal for the precise dermal segregation of CTS and DP fibroblasts and their elaboration of hair growth-inducing morphogens, DP fibroblasts could be tested for their hair growth-inducing properties in vivo,140,189 with and without the presence of NCAMblocking antibodies. Furthermore, the modulation of NCAM adhesiveness by the application of function-blocking anti-NCAM antibodies or PSA-digesting specific endosialidase65,66,190,191 in vivo and in organ culture of fetal or neonatal mouse skin could test whether the disruption of NCAM-mediated tissue integrity retards the condensation of DP/CTS fibroblasts and/or subsequent HF development. Also, since only a few “selector” genes have been reported to display binding sites for recognized MR molecules, further studies are urgently needed to link the striking hair phenotypes of published mouse mu-

Topobiology of the Hair Follicle

305

tants of potential “selector” genes such as Lef1174,176 to the morphoregulatory functions of specific adhesion molecules. It is likely that experiments of the kind suggested above will quickly help to answer many of the open key questions of trichotopobiology discussed earlier. The significance of rigorously addressing these questions reaches beyond the field of HF morphogenesis. On the one hand, the HF offers a fascinating, comparatively easily dissectable model microcosmos for studying the general role of adhesion forces in pattern formation and embryonic periodicity. On the other hand, given that the cyclic anagen development of mature HF rhythmically recapitulates central aspects of HF morphogenesis and that the capacity to induce anagen would be highly desirable for the treatment of many hair growth disorders associated with alopecia and/or effluvium,186 it is also clinically important to better understand the role of MR molecules in generating a new hair shaft factory, the anagen bulb. Thus, to better characterize the topobiology of the HF is an eminent challenge to developmental biologists, hair researchers, and clinicians alike.

Acknowledgments We thank Dr. Cheng-Ming Chuong for helpful critique and advice, and Carina van der Veen for assistance with the preparation of figures and tables. The compilation of this chapter and the authors’ quoted work was supported by grants from Deutsche Forschungsgemeinschaft to R. P. (Pa 345/3-2 and 8-1) and from Wella AG, Darmstadt.

References 1. Hardy MH. The secret life of the hair follicle. Trends Genet 1992; 8:55-61. 2. Danilenko DM, Ring BD, Pierce GF. Growth factors and cytokines in hair follicle development and cycling: Recent insights from animal models and the potentials for clinical therapy. Mol Med Today 1996b; 2:460-467. 3. Stenn KS, Combates NJ, Eilertsen KJ et al. Hair follicle growth controls. Dermatol Clinics 1996; 14:543-558. 4. Chuong CM, Chen HM, Jiang TX et al. Adhesion molecules in skin development: Morphogenesis of feather and hair. Ann N Y Acad Sci 1991; 642:263-280. 5. Chuong CM. The making of a feather: Homeoproteins, retinoids and adhesion molecules. BioEssays 1993; 15:513-521. 6. Chuong CM, Widelitz RB, Jiang TX. Adhesion molecules and homeoproteins in the phenotypic determination of skin appendages. J Invest Dermatol 1993; 101:10S-15S. 7. Edelman GM. Topobiology: An Introduction to Molecular Embryology. New York: Basic Books. 1988b. 8. Edelman GM. Morphoregulatory molecules. Biochemistry 1988a; 27:3533-3543. 9. Held IL. Models for Embryonic Periodicity Basel: Karger.1992. 10. Gilbert S. Developmental Biology. Sunderland, MA: Sinauer.1994. 11. Müller WF. Developmental Biology. New York: Springer. 1997. 12. Chen CS, Mrksich M, Huang S et al. Geometric control of life and death. Science 1997; 276:1425-1428. 13. Ruoslahti E. Stretching is good for a cell. Science 1997; 276:1345-1346. 14. Steinberg MS. Does differential adhesion govern self-assembly processes in histogenesis? Equilibrium configurations and the emergence of a hierarchy among populations of embryonic cells. J Exp Zool 1970; 173:395-433. 15. Martz E, Steinberg MS. The role of cell-cell contact in “contact” inhibition of cell division: A review and new evidence. J Cell Physiol 1972; 79:189-210. 16. Wiseman LL, Steinberg MS, Phillips HM. Experimental modulation of intercellular cohesiveness: reversal of tissue assembly patterns. Dev Biol 1972; 28:498-517. 17. Wiseman LL, Steinberg MS The movement of single cells within solid tissue masses. Exp Cell Res 1973; 79:468-471.

306

Molecular Basis of Epithelial Appendage Morphogenesis

18. Steinberg MS, Takeichi M. Experimental specification of cell sorting, tissue spreading, and specific spatial patterning by quantitative differences in cadherin expression. Proc Natl Acad Sci USA 1994; 91:206-209. 19. Lodish H, Baltimore D, Berk A et al. Molecular Cell Biology. New York: Scientific American Books. 1995. 20. Cooper GM. The Cell. Sunderland, MA: ASM Press/Sinauer, 1997. 21. Gumbiner BM. Cell adhesion: The molecular basis of tissue architecture and morphogenesis. Cell 1996; 84:345-357. 22. Huber O, Bierkamp C, Kemler R. Cadherins and catenins in development. Curr Opin Cell Biol 1996a; 8:685-691. 23. Kuhl M, Wedlich D. Xenopus cadherins: Sorting out types and functions in embryogenesis. Dev Dyn 1996; 207:121-134. 24. Redies C, Takeichi M. Cadherins in the developing central nervous system: An adhesive code for segmental and functional subdivisions. Dev Biol 1996; 180:413-423. 25. Suzuki ST. Protocadherins and diversity of the cadherin superfamily. J Cell Sci 1996; 109:2609-2611. 26. Takeichi M, Matsunami H, Inoue T et al. Roles of cadherins in patterning of the developing brain. Dev Neurosci 1997; 19:86-87. 27. Gumbiner BM. Epithelial morphogenesis. Cell 1992; 69:385-387. 28. Rodriguez Boulan E, Powell SK. Polarity of epithelial and neuronal cells. Annu Rev Cell Biol 1992; 8:395-427. 29. Rodriguez Boulan E, Zurzolo C. Polarity signals in epithelial cells. J Cell Sci Suppl 1993; 17:9-12. 30. Takeichi M, Watabe M, Shibamoto S et al. Cadherin-dependent organization and disorganization of epithelial architecture. Princess Takamatsu Symp 1994; 24:28-37. 31. Hermiston ML, Gordon JI. In vivo analysis of cadherin function in the mouse intestinal epithelium: essential roles in adhesion, maintenance of differentiation, and regulation of programmed cell death. J Cell Biol 1995; 129:489-506. 32. Le Gall AH, Yeaman C, Muesch A et al. Epithelial cell polarity: new perspectives. Semin Nephrol 1995; 15:272-284. 33. Ruoslahti E, Reed JC. Anchorage dependence, integrins, and apoptosis. Cell 1994; 77:477-478. 34. Frisch SM, Vuori K, Ruoslahti E et al. Control of adhesion-dependent cell survival by focal adhesion kinase. J Cell Biol 1996; 134:793-799. 35. Hermiston ML, Wong MH, Gordon JI. Forced expression of E-cadherin in the mouse intestinal epithelium slows cell migration and provides evidence for nonautonomous regulation of cell fate in a self-renewing system. Genes Dev 1996; 10:985-996. 36. Yamada KM, Geiger B. Molecular interactions in cell adhesion complexes. Curr Opin Cell Biol 1997; 9:76-85. 37. Kemler R, Ozawa M, Ringwald M. Calcium-dependent cell adhesion molecules. Curr Opin Cell Biol 1989; 1:892-897. 38. Nagafuchi A, Shirayoshi Y, Okazaki K et al. Transformation of cell adhesion properties by exogenously introduced E-cadherin cDNA. Nature 1987; 329:341-343. 39. McNeill H, Ozawa M, Kemler R et al. Novel function of the cell adhesion molecule uvomorulin as an inducer of cell surface polarity. Cell 1990; 62:309-316. 40. Gumbiner B, Stevenson B, Grimaldi A. The role of the cell adhesion molecule uvomorulin in the formation and maintenance of the epithelial junctional complex. J Cell Biol 1988; 107:1575-1587. 41. Jongen WM, Fitzgerald DJ, Asamoto M et al. Regulation of connexin 43-mediated gap junctional intercellular communication by Ca2+ in mouse epidermal cells is controlled by E-cadherin. J Cell Biol 1991; 114:545-555. 42. Watabe M, Nagafuchi A, Tsukita S et al. Induction of polarized cell-cell association and retardation of growth by activation of the E-cadherin-catenin adhesion system in a dispersed carcinoma line. J Cell Biol 1994; 127:247-256.

Topobiology of the Hair Follicle

307

43. Tang A, Amagai M, Granger LG et al. Adhesion of epidermal Langerhans cells to keratinocytes mediated by E-cadherin. Nature 1993; 361:82-85. 44. Cepek KL, Shaw SK, Parker CM et al. Adhesion between epithelial cells and T lymphocytes mediated by E-cadherin and the alpha E beta 7 integrin. Nature 1994; 372:190-193. 45. Karecla PI, Bowden SJ, Green SJ et al. Recognition of E-cadherin on epithelial cells by the mucosal T cell integrin alpha M290 beta 7 alpha E beta 7. Eur J Immunol 1995; 25:852-856. 46. Behrens J, von Kries JP, Kuhl M et al. Functional interaction of beta-catenin with the transcription factor Lef1. Nature 1996; 382:638-642. 47. Huber O, Korn R, McLaughlin J. Nuclear localization of beta-catenin by interaction with transcription factor Lef1. Mech Dev 1996b; 59:3-10. 48. Moles JP, Watt FM. The epidermal stem cell compartment: variation in expression levels of E-cadherin and catenins within the basal layer of human epidermis. J Histochem Cytochem 1997; 457:867-874. 49. Nose A, Takeichi M. A novel cadherin cell adhesion molecule: Its expression patterns associated with implantation and organogenesis of mouse embryos. J Cell Biol 1986; 103:2649-2658. 50. Fujita M, Furukawa F, Fujii K et al. Expression of cadherin cell adhesion molecules during human skin development: Morphogenesis of epidermis, hair follicles and eccrine sweat ducts. Arch Dermatol Res 1992; 284:159-166. 51. Springer TA. Adhesion receptors of the immune system. Nature 1990; 346:425-434. 52. Baldwin TJ, Fazeli MS, Doherty P et al. Elucidation of the molecular actions of NCAM and structurally related cell adhesion molecules. J Cell Biochem 1996; 61:502-513. 53. Brummendorf T, Rathjen FG. Structure/function relationships of axon-associated adhesion receptors of the immunoglobulin superfamily. Curr Opin Neurobiol 1996; 6:584-593. 54. Kelm S, Schauer R, Crocker PR. The Sialoadhesins—a family of sialic acid-dependent cellular recognition molecules within the immunoglobulin superfamily. Glycoconj J 1996; 13:913-926. 55. Rutishauser U. Adhesion molecules of the nervous system. Curr Opin Neurobiol 1993; 3:709-715. 56. Cunningham BA, Hemperly JJ, Murray BA et al. Neural cell adhesion molecule: Structure, immunoglobulin-like domains, cell surface modulation, and alternative RNA splicing. Science 1987; 236:799-806. 57. Santoni MJ, Barthels D, Barbas JA et al. Analysis of cDNA clones that code for the transmembrane forms of the mouse neural cell adhesion molecule NCAM and are generated by alternative RNA splicing. Nucleic Acids Res 1987; 15:8621-8641. 58. Small SJ, Haines SL, Akeson RA. Polypeptide variation in an N-CAM extracellular immunoglobulin-like fold is developmentally regulated through alternative splicing. Neuron 1988; 1:1007-1017. 59. Thompson J, Dickson G, Moore SE et al. Alternative splicing of the neural cell adhesion molecule gene generates variant extracellular domain structure in skeletal muscle and brain. Genes Dev 1989; 3:348-357. 60. Gennarini G, Hirn M, Deagostini Bazin H et al. Studies on the transmembrane disposition of the neural cell adhesion molecule N-CAM. The use of liposome-inserted radioiodinated N-CAM to study its transbilayer orientation. Eur J Biochem 1984a; 142:65-73. 61. Gennarini G, Rougon G, Deagostini Bazin H et al. Studies on the transmembrane disposition of the neural cell adhesion molecule N-CAM. A monoclonal antibody recognizing a cytoplasmic domain and evidence for the presence of phosphoserine residues. Eur J Biochem 1984b; 142:57-64. 62. Rutishauser U, Acheson A, Hall AK et al. The neural cell adhesion molecule NCAM as a regulator of cell-cell interactions. Science 1988; 240:53-57. 63. Pollerberg EG, Sadoul R, Goridis C et al. Selective expression of the 180-kD component of the neural cell adhesion molecule N-CAM during development. J Cell Biol 1985; 101:1921-1929.

308

Molecular Basis of Epithelial Appendage Morphogenesis

64. Murray BA, Hemperly JJ, Prediger EA et al. Alternatively spliced mRNAs code for different polypeptide chains of the chicken neural cell adhesion molecule N-CAM. J Cell Biol 1986; 102:189-193. 65. Acheson A, Sunshine JL, Rutishauser U. NCAM polysialic acid can regulate both cell-cell and cell-substrate interactions. J Cell Biol 1991; 114:143-153. 66. Ono K, Tomasiewicz H, Magnuson T et al. N-CAM mutation inhibits tangential neuronal migration and is phenocopied by enzymatic removal of polysialic acid. Neuron 1994; 13:595-609. 67. Edelman GM. Cell adhesion molecules. Science 1983; 219:450-457. 68. Chuong CM, Edelman GM. Expression of cell-adhesion molecules in embryonic induction. II. Morphogenesis of adult feathers. J Cell Biol 1985; 101:1027-1043. 69. Crossin KL, Chuong CM, Edelman GM. Expression sequences of cell adhesion molecules. Proc Natl Acad Sci USA 1985; 82:6942-6946. 70. Edelman GM, Jones FS. Developmental control of N-CAM expression by Hox and Pax gene products. Philos Trans R Soc Lond B Biol Sci 1995; 349:305-312. 71. Brackenbury R. Expression of neural cell adhesion molecules in normal and pathologic tissues. Ann N Y Acad Sci 1988; 540:39-46. 72. Martini R, Schachner M. Immunoelectron microscopic localization of neural cell adhesion molecules L1, N-CAM, and myelin-associated glycoprotein in regenerating adult mouse sciatic nerve. J Cell Biol 1988; 106:1735-1746. 73. Martini R. Expression and functional roles of neural cell surface molecules and extracellular matrix components during development and regeneration of peripheral nerves. J Neurocytol 1994; 23:1-28. 74. Botchkarev VA, Eichmüller S, Johansson O et al. Hair cycle-dependent plasticity of skin and hair follicle innervation in normal murine skin. J Comp Neurol, 1997; 386(3):379-395. 75. Reichardt LF, Tomaselli KJ. Extracellular matrix molecules and their receptors: Functions in neural development. Annu Rev Neurosci 1991; 14:531-570. 76. Roubin R, Deagostini Bazin H, Hirsch MR et al. Modulation of NCAM expression by transforming growth factor-beta, serum, and autocrine factors. J Cell Biol 1990; 111:673-684. 77. Jiang TX, Yi JR, Ying SY et al. Activin enhances chondrogenesis of limb bud cells: Stimulation of precartilaginous mesenchymal condensations and expression of NCAM. Dev Biol 1993; 155:545-557. 78. Einheber S, Hannocks MJ, Metz CN et al. Transforming growth factor-beta 1 regulates axon/Schwann cell interactions. J Cell Biol 1995; 129:443-458. 79. Stewart HJ, Rougon G, Dong Z et al. TGF-betas upregulate NCAM and L1 expression in cultured Schwann cells, suppress cyclic AMP-induced expression of O4 and galactocerebroside, and are widely expressed in cells of the Schwann cell lineage in vivo. Glia 1995; 15:419-436. 80. Ting Berreth SA, Chuong CM. Local delivery of TGF beta2 can substitute for placode epithelium to induce mesenchymal condensation during skin appendage morphogenesis. Dev Biol 1996; 179:347-359. 81. Jones FS, Holst BD, Minowa O et al. Binding and transcriptional activation of the promoter for the neural cell adhesion molecule by HoxC6 Hox-3.3. Proc Natl Acad Sci USA 1993; 90:6557-6561. 82. Valarche I, Tissier Seta JP et al. The mouse homeodomain protein Phox2 regulates Ncam promoter activity in concert with Cux/CDP and is a putative determinant of neurotransmitter phenotype. Development 1 1993; 19:881-896. 83. Holst BD, Goomer RS, Wood IC et al. Binding and activation of the promoter for the neural cell adhesion molecule by Pax-8. J Biol Chem 1994; 269:22245-22252. 84. Edelman GM. Morphoregulation. Dev Dyn 1992;193:2-10. 85. Holst BD, Wang Y, Jones FS et al. A binding site for Pax proteins regulates expression of the gene for the neural cell adhesion molecule in the embryonic spinal cord. Proc Natl Acad Sci USA 1997; 94:1465-1470. 86. Varner JA, Cheresh DA. Integrins and cancer. Curr Opin Cell Biol 1996; 8:724-730.

Topobiology of the Hair Follicle

309

87. Hynes RO. Integrins: versatility, modulation, and signaling in cell adhesion. Cell 1992; 69:11-25. 88. Cardarelli PM, Cobb RR, Nowlin DM et al. Cyclic RGD peptide inhibits alpha 4 beta 1 interaction with connecting segment 1 and vascular cell adhesion molecule. J Biol Chem 1994; 269:18668-18673. 89. Hirasawa M, Shijubo N, Uede T et al. Integrin expression and ability to adhere to extracellular matrix proteins and endothelial cells in human lung cancer lines. Br J Cancer 1994; 70:466-473. 90. Kjoller I, Kanse SM, Kirkegaard T et al. Plasminogen activator inhibitor-1 represses integrinand vitronectin-mediated cell migration independently of its function as an inhibitor of plasminogen activation. Exp Cell Res 1997; 232:420-429. 91. Fässler R, Georges Labouesse E, Hirsch E. Genetic analyses of integrin function in mice. Curr Opin Cell Biol 1996; 8:641-646. 92. Wang N, Butler JP, Ingber DE. Mechanotransduction across the cell surface and through the cytoskeleton. Science 1993; 260:1124-1127. 93. Perris R. The extracellular matrix in neural crest-cell migration. Trends Neurosci 1997; 20:23-31. 94. Lafrenie RM, Yamada KM. Integrin-dependent signal transduction. J Cell Biochem 1996; 61:543-553. 95. Hanks SK, Polte TR. Signaling through focal adhesion kinase. BioEssays 1997; 19:137-145. 96. Roman J. Fibronectin and fibronectin receptors in lung development. Exp Lung Res 1997; 23:147-159. 97. Sjaastad MD, Nelson WJ. Integrin-mediated calcium signaling and regulation of cell adhesion by intracellular calcium. BioEssays 1997; 19:47-55. 98. Todd RF, Petty HR. Beta 2 CD11/CD18 integrins can serve as signaling partners for other leukocyte receptors. J Lab Clin Med 1997; 129:492-498. 99. Wei J, Shaw LM, Mercurio AM. Integrin signaling in leukocytes: Lessons from the alpha6beta1 integrin. J Leukoc Biol 1997; 61:397-407. 100. Yoshimura Y. Integrins: expression, modulation, and signaling in fertilization, embryogenesis and implantation. Keio J Med 1997; 46:16-24. 101. Schwartz MA, Both G, Lechene C. Effect of cell spreading on cytoplasmic pH in normal and transformed fibroblasts. Proc Natl Acad Sci USA 1989; 86:4525-4529. 102. Schwartz MA, Cragoe EJ Jr, Lechene CP. pH regulation in spread cells and round cells. J Biol Chem 1990; 265:1327-1332. 103. Schwartz MA, Lechene C, Ingber DE. Insoluble fibronectin activates the Na/H antiporter by clustering and immobilizing integrin alpha 5 beta 1, independent of cell shape. Proc Natl Acad Sci USA 1991; 88:7849-7853. 104. Pelletier AJ, Bodary SC, Levinson AD. Signal transduction by the platelet integrin alpha IIb beta 3: Induction of calcium oscillations required for protein-tyrosine phosphorylation and ligand-induced spreading of stably transfected cells. Mol Biol Cell 1992; 3:989-998. 105. Leavesley DI, Schwartz MA, Rosenfeld M et al. Integrin beta 1- and beta 3-mediated endothelial cell migration is triggered through distinct signaling mechanisms. J Cell Biol 1993; 121:163-170. 106. Schwartz MA. Spreading of human endothelial cells on fibronectin or vitronectin triggers elevation of intracellular free calcium. J Cell Biol 1993; 120:1003-1010. 107. McNamee HP, Ingber DE, Schwartz MA. Adhesion to fibronectin stimulates inositol lipid synthesis and enhances PDGF-induced inositol lipid breakdown. J Cell Biol 1993; 121:673-678. 108. Guadagno TM, Ohtsubo M, Roberts JM et al. A link between cyclin A expression and adhesion-dependent cell cycle progression. Science 1993; 262:1572-1575. 109. Rich S, Van Nood N, Lee HM. Role of alpha 5 beta 1 integrin in TGF-beta 1-costimulated CD8+ T cell growth and apoptosis. J Immunol 1996; 157:2916-2923. 110. Ginis I, Faller DV. Protection from apoptosis in human neutrophils is determined by the surface of adhesion. Am J Physiol 1997; 272:C295-309.

310

Molecular Basis of Epithelial Appendage Morphogenesis

111. Crouch DH, Fincham VJ, Frame MC. Targeted proteolysis of the focal adhesion kinase pp125 FAK during c-MYC-induced apoptosis is suppressed by integrin signaling. Oncogene 1996; 12:2689-2696. 112. Scott G, Cassidy L, Busacco A. Fibronectin suppresses apoptosis in normal human melanocytes through an integrin-dependent mechanism. J Invest Dermatol 1997; 108:147-153. 113. Weedon D, Strutton G. Apoptosis as the mechanism of involution of hair follicles in catagen transformation. Acta Derm Venereol 1981; 61:335-339. 114. Paus R, Rosenbach T, Haas N et al. Patterns of cell death: The significance of apoptosis for dermatology. Exp Dermatol 1993; 2:3-11. 115. Polakowska RR, Haake A. Apoptosis: The skin from a new perspective. Death Differentiation 1994; 1:19-31. 116. Lindner G, Botchkarev VA, Botchkarev NV et al. Analysis of apoptosis during hair follicle regression. Am J Pathol 1997; 151:1601-1617. 117. Paus R, Handjiski B, Czarnetzki BM et al. Biologie des Haarfollikels. Hautarzt 1994a; 45:808-825. 118. Paus R. Control of the hair cycle and hair diseases as cycling disorders. Curr Opin Dermatol 1996; 3:248-258. 119. Paus R, Foitzik K, Welker P et al. Transforming growth factor-β receptor type I and type II expression during murine hair follicle development and cycling. J Invest Dermatol 1997; 109:518-526. 120. Montagna W, Ellis RA. The biology of hair growth. New York: Academic Press.1958. 121. Stenn KS, Messenger AG, Baden HP. The molecular and structural biology of hair. Ann N Y Acad Sci 1991; 642:1-519. 122. Hirai Y, Nose A, Kobayashi S et al. Expression and role of E- and P-cadherin adhesion molecules in embryonic histogenesis. II. Skin morphogenesis. Development 1989; 105:271-277. 123. Müller-Röver S, Tokura Y, Paus R. Topobiology of the Hair Follicle: I. Expression of CellAdhesion Molecules During Neonatal Hair Follicle Morphogenesis in mice. J Invest Dermatol 1996; 107:654a. 124. Müller-Röver S, Tokura Y, Paus R et al. E- and P-cadherin expression during murine hair follicle development and cycling. Exp Dermatol in press. 125. Kaplan ED, Holbrook KA. Dynamic expression patterns of tenascin, proteoglycans, and cell adhesion molecules during human hair follicle morphogenesis. Dev Dyn 1994; 199:141-155. 126. Mege RM, Matsuzaki F, Gallin WJ et al. Construction of epithelioid sheets by transfection of mouse sarcoma cells with cDNAs for chicken cell adhesion molecules. Proc Natl Acad Sci USA 1988; 85:7274-7278. 127. Cotsarelis G, Sun TT, Lavker RM. Label-retaining cells reside in the bulge area of pilosebaceous unit: Implications for follicular stem cells, hair cycle, and skin carcinogenesis. Cell 1990; 61:1329-1337. 128. Sun TT, Cotsarelis G, Lavker RM. Hair follicular stem cells: The bulge-activation hypothesis. J Invest Dermatol 1991; 96:77S-78S. 129. Lavker RM, Sun TT. Hair follicle stem cells: Present concepts. J Invest Dermatol 1995; 104:38S-39S. 130. Wilson C, Cotsarelis G, Wei ZG et al. Cells within the bulge region of mouse hair follicle transiently proliferate during early anagen: Heterogeneity and functional differences of various hair cycles. Differentiation 1994; 55:127-136. 131. Hodivala KJ, Watt FM. Evidence that cadherins play a role in the downregulation of integrin expression that occurs during keratinocyte terminal differentiation. J Cell Biol 1994; 124:589-600. 132. Zhu AJ, Watt FM. Expression of a dominant negative cadherin mutant inhibits proliferation and stimulates terminal differentiation of human epidermal keratinocytes. J Cell Sci 1996; 109:3013-3023.

Topobiology of the Hair Follicle

311

133. Gallin WJ, Chuong CM, Finkel LH et al. Antibodies to liver cell adhesion molecule perturb inductive interactions and alter feather pattern and structure. Proc Natl Acad Sci USA 1986; 83:8235-8239. 134. Botchkarev VA, Paus R, Czarnetzki BM et al. Hair cycle-dependent changes in mast cell histochemistry in murine skin. Arch Dermatol Res 1995; 287:683-686. 135. Vielkind U, Sebzda MK, Gibson IR et al. Dynamics of Merkel cell patterns in developing hair follicles in the dorsal skin of mice, demonstrated by a monoclonal antibody to mouse keratin 8. Acta Anat 1995; 152:93-109. 136. Jiang TX, Chuong CM. Mechanism of skin morphogenesis. I. Analyses with antibodies to adhesion molecules tenascin, N-CAM, and integrin. Dev Biol 1992; 150:82-98. 137. Müller-Röver S, Botchkarev V, Panteleyev A et al. NCAM expression during hair follicle morphogenesis and cycling. J Invest Dermatol 1997a; 108:654a. 138. Müller-Röver S, van der Veen C, Panteleyev A et al. Distinct patterns of NCAM expression are associated with defined stages of murine hair follicle morphogenesis and cycling. 1997c; submitted. 139. Jahoda CA, Horne KA, Oliver RF. Induction of hair growth by implantation of cultured dermal papilla cells. Nature 1984; 311:560-562. 140. Jahoda AB, Reynolds AJ. Dermal-Epidermal Interactions—Adult Follicle-Derived Cell Populations and Hair Growth. Dermatologic Clinics 1996; 14:573-583. 141. Reynolds AJ, Jahoda CA. Hair matrix germinative epidermal cells confer follicle-inducing capabilities on dermal sheath and high passage papilla cells. Development 1996; 122:3085-3094. 142. Saffell JL, Williams EJ, Mason IJ et al. Expression of a dominant negative FGF receptor inhibits axonal growth and FGF receptor phosphorylation stimulated by CAMs. Neuron 1997; 18:231-242. 143. Kreidberg JA, Donovan MJ, Goldstein SL et al. Alpha 3 beta 1 integrin has a crucial role in kidney and lung organogenesis. Development 1996; 122:3537-3547. 144. Yang JT, Rayburn H, Hynes RO. Cell adhesion events mediated by alpha 4 integrins are essential in placental and cardiac development. Development 1995; 121:549-560. 145. Sundberg JP. Handbook of mouse mutations with skin and hair abnormalities. Boca Roton: CRC press. 1994. 146. Botchkarev VA, Lewin GA, Albers KM et al. Neurotrophins and murine hair follicle morphogenesis: Expression patterns of NT-3, NT-4, BDNF, Trk B, Trk C and indications for a functional role in hair follicle regression and cycling. J Invest Dermatol 1997b; 108:620a. 147. Maurer M, Fische E, Handjiski B et al. Activated skin mast cells are involved in hair follicle regression catagen 9. Lab Invest 1997a; in press. 148. Wang A, Patrone L, McDonald JA et al. Expression of the integrin subunit alpha 9 in the murine embryo. Dev Dyn 1995; 204:421-431. 149. Huang XZ, Wu JF, Cass D et al. Inactivation of the integrin beta 6 subunit gene reveals a role of epithelial integrins in regulating inflammation in the lung and skin. J Cell Biol 1996; 133:921-928. 150. Carroll JM, Romero MR, Watt FM. Suprabasal integrin expression in the epidermis of transgenic mice results in developmental defects and a phenotype resembling psoriasis. Cell 1995; 83:957-968. 151. Morgan BA. Hox genes and embryonic development. Poult Sci 1997; 76:96-104. 152. Sharkey M, Graba Y, Scott MP. Hox genes in evolution: Protein surfaces and paralog groups. Trends Genet 1997; 13:145-151. 153. Lewin B. Genes VI. New York: Oxford University Press. 1997. 154. Lufkin T. Transcriptional control of Hox genes in the vertebrate nervous system. Curr Opin Genet Dev 1996; 6:575-580. 155. Marshall H, Morrison A, Studer M et al. Retinoids and Hox genes. Faseb J 1996; 10:969-978. 156. Mundlos S, Olsen BR. Heritable diseases of the skeleton. Part I: Molecular insights into skeletal development-transcription factors and signaling pathways. Faseb J 1997; 11:125-132. 157. Chen F, Capecchi MR. Targeted mutations in hoxa-9 and hoxb-9 reveal synergistic interactions. Dev Biol 1997; 181:186-196.

312

Molecular Basis of Epithelial Appendage Morphogenesis

158. McGinnis W, Krumlauf R. Homeobox genes and axial patterning. Cell 1992; 68:283-302. 159. Chuong CM, Oliver G, Ting SA et al. Gradients of homeoproteins in developing feather buds. Development 1990; 110:1021-1030. 160. Bieberich CJ, Ruddle FH, Stenn KS. Differential expression of the Hox 3.1 gene in adult mouse skin. Ann N Y Acad Sci 1991; 642:346-353. 161. Brown WM, Stenn KS. Homeobox genes and the patterning of skin diseases. J Cutan Pathol 1993; 20:289-293. 162. Detmer K, Lawrence HJ, Largman C. Expression of class I homeobox genes in fetal and adult murine skin. J Invest Dermatol 1993; 101:517-522. 163. Mathews CH, Detmer K, Lawrence HJ et al. Expression of the Hox 2.2 homeobox gene in murine embryonic epidermis. Differentiation 1993; 52:177-184. 164. Scott GA, Goldsmith LA. Homeobox genes and skin development: A review. J Invest Dermatol 1993; 101:3-8. 165. Brown WM, Zhou L, Taylor GR. The nucleotide sequence of the murine Hox-D3 (Hox4.1) gene reveals extensive identity with the human protein. Biochim Biophys Acta 1994; 1219:219-222. 166. Chalepakis G, Wijnholds J, Giese P et al. Characterization of Pax-6 and Hoxa-1 binding to the promoter region of the neural cell adhesion molecule L1. DNA Cell Biol 1994; 13:891-900. 167. Rieger E, Bijl JJ, van Oostveen JW et al. Expression of the homeobox gene HoxC4 in keratinocytes of normal skin and epithelial skin tumors is correlated with differentiation. J Invest Dermatol 1994; 103:341-346. 168. Noveen A, Jiang TX, Ting Berreth SA et al. Homeobox genes Msx-1 and Msx-2 are associated with induction and growth of skin appendages. J Invest Dermatol 1995; 104:711-719. 169. Chuong CM, Widelitz RB, Ting Berreth S et al. Early events during avian skin appendage regeneration: dependence on epithelial-mesenchymal interaction and order of molecular reappearance. J Invest Dermatol 1996; 107:639-646. 170. Lamb TM, Harland RM. Fibroblast growth factor is a direct neural inducer, which combined with noggin generates anterior-posterior neural pattern. Development 1995; 121:3627-3636. 171. Kostakopoulou K, Vogel A, Brickell P et al. ‘Regeneration’ of wing bud stumps of chick embryos and reactivation of Msx-1 and Shh expression in response to FGF-4 and ridge signals. Mech Dev 1996; 55:119-131. 172. Pownall ME, Tucker AS, Slack JM et al. eFgf, Xcad3 and Hox genes form a molecular pathway that establishes the anteroposterior axis in Xenopus. Development 1996; 122:3881-3892. 173. Cohn MJ, Patel K, Krumlauf R et al. Hox9 genes and vertebrate limb specification. Nature 1997; 387:97-101. 174. Zhou P, Byrne C, Jacobs J et al. Lymphoid enhancer factor 1 directs hair follicle patterning and epithelial cell fate. Genes Dev 1995; 9:700-713. 175. Kratochwil K, Dull M, Farinas I et al. Lef1 expression is activated by BMP-4 and regulates inductive tissue interactions in tooth and hair development. Genes Dev 1996; 10:1382-1394. 176. van Genderen C, Okamura RM, Farinas I et al. Development of several organs that require inductive epithelial-mesenchymal interactions is impaired in Lef1-deficient mice. Genes Dev 1994; 8:2691-2703. 177. Danilenko DM, Ring BD, Pierce GF. Growth factors and cytokines in hair follicle development and cycling: Recent insights from animal models and the potentials for clinical therapy. Mol Med Today 1996a; 2:460-467. 178. Tomasiewicz H, Ono K, Yee D et al. Genetic deletion of a neural cell adhesion molecule variant N-CAM-180 produces distinct defects in the central nervous system. Neuron 1993; 11:1163-1174. 179. Cremer H, Lange R, Christoph A et al. Inactivation of the N-CAM gene in mice results in size reduction of the olfactory bulb and deficits in spatial learning. Nature 1994; 367:455-459.

Topobiology of the Hair Follicle

313

180. Bullard DC, Scharffetter Kochanek K, McArthur MJ et al. A polygenic mouse model of psoriasiform skin disease in CD18-deficient mice. Proc Natl Acad Sci USA 1996; 93:2116-2121. 181. Schmits R, Kundig TM, Baker DM et al. LFA-1-deficient mice show normal CTL responses to virus but fail to reject immunogenic tumor. J Exp Med 1996; 183:1415-1426. 182. Lu H, Smith CW, Perrard J et al. LFA-1 is sufficient in mediating neutrophil emigration in Mac-1-deficient mice. J Clin Invest 1997; 99:1340-1350. 183. Fässler R, Meyer M. Consequences of lack of beta 1 integrin gene expression in mice. Genes Dev 1995; 9:1896-1908. 184. Li L, Paus R, Slominski A et al. Skin histoculture assay for studying the hair cycle [letter]. In Vitro Cell Dev Biol 1992; 28A:695-698. 185. Paus R, Handjiski B, Czarnetzki BM et al. A murine model for inducing and manipulating hair follicle regression catagen: Effects of dexamethasone and cyclosporin A. J Invest Dermatol 1994b; 103:143-147. 186. Paus R, Handjiski B, Eichmuller S et al. Chemotherapy-induced alopecia in mice. Induction by cyclophosphamide, inhibition by cyclosporine A, and modulation by dexamethasone. Am J Pathol 1994c; 144:719-734. 187. Paus R, Schilli MB, Handjiski B et al. Topical calcitriol enhances normal hair regrowth but does not prevent chemotherapy-induced alopecia in mice. Cancer Res 1996; 56:4438-4443. 188. Maurer M, Handjiski B, Paus R. Hair growth modulation by topical immunophilin ligands: Induction of anagen, inhibition of massive catagen development, and relative protection from chemotherapy-induced alopecia. Am J Pathol 1997b; 150:1433-1441. 189. Williams IR, Kupper TS. Epidermal expression of intercellular adhesion molecule 1 is not a primary inducer of cutaneous inflammation in transgenic mice. Proc Natl Acad Sci USA 1994; 91:9710-9714. 190. Jahoda CA, Oliver RF. Vibrissa dermal papilla cell aggregative behaviour in vivo and in vitro. J Embryol Exp Morphol 1984; 79:211-224. 191. Landmesser L, Dahm L, Tang JC et al. Polysialic acid as a regulator of intramuscular nerve branching during embryonic development. Neuron 1990; 4:655-667. 192. Becker CG, Artola A, Gerardy Schahn R et al. The polysialic acid modification of the neural cell adhesion molecule is involved in spatial learning and hippocampal long-term potentiation. J Neurosci Res 1996; 45:143-152. 193. Hynes RO, Yamada KM. Fibronectins: Multifunctional modular glycoproteins. J Cell Biol 1982; 95:369-377. 194. Hynes RO. Integrins: A family of cell surface receptors. Cell 1987; 48:549-554. 195. Salmivirta K, Gullberg D, Hirsch E et al. Integrin subunit expression associated with epithelial-mesenchymal interactions during murine tooth development. Dev Dyn 1996; 205:104-113. 196. Stephens LE, Sutherland AE, Klimanskaya IV et al. Deletion of beta 1 integrins in mice results in inner cell mass failure and peri-implantation lethality. Genes Dev 1995; 9:1883-1895. 197. Paus R, Link RE. The psoriatic epidermal lesion and anagen hair growth may share the same “switch-on” mechanism. Yale J Biol Med 1988; 61:467-476. 198. Albelda SM, Buck CA. Integrins and other cell adhesion molecules. Faseb J 1990; 4:2868-2880. 199. Van der Vieren M, Le Trong H, Wood CL et al. A novel leukointegrin, alpha d beta 2, binds preferentially to ICAM-3. Immunity 1995; 3:683-690. 200. Larson RS, Springer TA. Structure and function of leukocyte integrins. Immunol Rev 1990; 114:181-217. 201. Pilewski JM, Albelda SM. Cell adhesion molecules in asthma: Homing, activation, and airway remodeling. Am J Respir Cell Mol Biol 1995; 12:1-3. 202. Springer TA. Traffic signals on endothelium for lymphocyte recirculation and leukocyte emigration. Annu Rev Physiol 1995; 57:827-872. 203. Hynes RO, Wagner DD. Genetic manipulation of vascular adhesion molecules in mice. J Clin Invest 1996; 98:2193-2195.

314

Molecular Basis of Epithelial Appendage Morphogenesis

204. van der Neut R, Krimpenfort P, Calafat J et al. Epithelial detachment due to absence of hemidesmosomes in integrin beta 4 null mice. Nat Genet 1996; 13:366-369. 205. Yamada S, Brown KE, Yamada KM. Differential mRNA regulation of integrin subunits alpha V, beta 1, beta 3, and beta 5 during mouse embryonic organogenesis. Cell Adhes Commun 1995; 3:311-325. 206. Yamada S, Yamada KM, Brown KE. Integrin regulatory switching in development: oscillation of beta 5 integrin mRNA expression during epithelial-mesenchymal interactions in tooth development. Int J Dev Biol 1994; 38:553-556. 207. Breuss JM, Gallo J, DeLisser HM et al. Expression of the beta 6 integrin subunit in development, neoplasia and tissue repair suggests a role in epithelial remodeling. J Cell Sci 1995; 108:2241-2251. 208. Yang JT, Rayburn H, Hynes RO. Embryonic mesodermal defects in alpha 5 integrin-deficient mice. Development 1993; 119:1093-1105. 209. Sonnenberg A, Calafat J, Janssen H et al. Integrin alpha 6/beta 4 complex is located in hemidesmosomes, suggesting a major role in epidermal cell-basement membrane adhesion. J Cell Biol 1991; 113:907-917. 210. Stepp MA, Spurr Michaud S et al. Alpha 6 beta 4 integrin heterodimer is a component of hemidesmosomes. Proc Natl Acad Sci USA 1990; 87:8970-8974. 211. Sutherland AE, Calarco PG, Damsky CH. Developmental regulation of integrin expression at the time of implantation in the mouse embryo. Development 1993; 119:1175-1186. 212. Damsky C, Sutherland A, Fisher S. Extracellular matrix 5: Adhesive interactions in early mammalian embryogenesis, implantation, and placentation. Faseb J 1993; 7:1320-1329. 213. Sorokin L, Sonnenberg A, Aumailley M et al. Recognition of the laminin E8 cell-binding site by an integrin possessing the alpha 6 subunit is essential for epithelial polarization in developing kidney tubules. J Cell Biol 1990; 111:1265-1273. 214. Georges-Labouesse E, Messaddeq N, Yehia G et al. Absence of integrin alpha 6 leads to epidermolysis bullosa and neonatal death in mice. Nat Genet 1996; 13:370-373. 215. Peltonen J, Larjava H, Jaakkola S et al. Localization of integrin receptors for fibronectin, collagen, and laminin in human skin. Variable expression in basal and squamous cell carcinomas. J Clin Invest 1989; 84:1916-1923.

CHAPTER 16

Late Events and the Regulation of Keratinocyte Differentiation in Hair and Feather Follicles George E. Rogers, Stephanie Dunn and Barry Powell

T

he late events of keratinocyte differentiation discussed here are mainly those that occur in the hair follicle. The reason for this emphasis is that more is known about the keratinization process of hair follicles than of feather follicles. The hair shaft keratinocyte is dedicated to the rapid synthesis of at least 70 different hair keratins. There are two groups of proteins, the keratin intermediate filament proteins and the keratin associated proteins, some of which combine to produce strikingly ordered arrays of filaments and matrix in the hair cortex whereas others produce amorphous structures in the hair cuticle. Near the end of hair differentiation the oxidation of the keratin proteins occurs to form a highly crosslinked, insoluble structure. Surrounding the hair shaft, the inner root sheath and outer root sheath layers provide biochemical and physical support to the growing hair. In the center of many hair fibers there is a column of medullary cells which provides stiffness to the fiber and promotes the insulating properties of hairs. The inner root sheath, adjacent to the growing hair, undergoes a hardening process involving the protein trichohyalin and two key enzymes, peptidylarginine deiminase and transglutaminase. Trichohyalin is also produced in great amounts in the medulla, where its fate is quite different. The inner root sheath does not normally emerge with the hair and is believed to be degraded by the action of the sebaceous gland. Many of the genes for the structural proteins of hair have been characterized and their patterns of expression mapped. The keratin intermediate filament genes are the first to express in the upper region of the follicle bulb, followed by the families of keratin associated genes in various cell-specific and stage-specific patterns. Our understanding of the genetic control of hair keratin gene expression is just beginning, but one transcription factor, lymphoid enhancer factor 1, already appears to have a central role in the regulation of many hair genes, with the ability to coordinate the interactions of other regulatory factors. Other possible regulatory elements have been identified, but the majority of them remain to be examined for functionality. The hair keratin intermediate filament proteins are related to the epidermal keratin intermediate filament proteins in what appears to be a shared evolutionary theme in the differentiation of the epidermis and its appendages. Furthermore, in the hair follicle both epidermal and hair types of keratins are expressed, but in separate compartments (outer root sheath for the epidermal type). Similarly, the filiform papillae of the tongue appear to

Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

316

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 16.1. Diagrammatic representation of the main layers of an adult hair follicle. The zones where important events of hair growth occur are indicated. It should be noted that epithelial cells of the bulb immediately surrounding the dermal papilla are derived from dividing cells in the base of the bulb. Some proliferation takes place at the upper level as well but at a much lower rate. Modified from Powell BC et al. In: Jolles P et al, eds. Formation and Structure of Hair. Basel: Birkhauser Verlag, 1997:59-148; reprinted with permission.

express both types in separate locations whereas in nail, although there is separate expression in cellular compartments, there is also evidence for coexpression of both types. As in the hair follicle, keratinization in the feather follicle is also a rapid process, and depends on the expression of a tandem array of many genes which encode small homologous proteins called β-keratins. Moreover, in an analogous manner the avian feather keratins are evolutionarily related to the β-keratins of scale, beak and claw. Despite the name, feather keratin proteins bear no structural relationship to the hair keratin proteins, and in the embryonic

Late Events and Regulation of Keratinocyte Differentiation in Hair and Feather Follicles

317

feather these proteins aggregate to produce specific 4 nm filaments. There is evidence that a regulatory element located in an intron controls feather keratin gene expression, but little else is known.

Introduction Once the hair follicle has formed, synthesis of the hair shaft requires the activation of a multitude of genes encoding hair structural proteins. The production of hair keratin mRNAs begins in the upper region of the follicle bulb as cells cease dividing and commit to terminal differentiation. Sequential activation of structural genes for keratin associated proteins follows, and a period of rapid protein synthesis ensues, until the cells have migrated approximately a third of the way up the follicle. Terminal differentiation is completed in two to three days, at which stage the hardened hair fiber has formed. This process of hair formation continues until the onset of the catagen phase of the hair cycle, when cell proliferation ceases and the follicle essentially shuts down. These late events in hair follicle differentiation occur rapidly and the hair-forming keratinocytes are structured for synthesis of the keratin protein product that remains in the cell. They have a cytoplasm packed with ribosomes, very little or no endoplasmic reticulum, few mitochondria and a relatively large nucleus.

The Proteins and Structures Formed in the Late Events of Hair Keratinization The product of the hair follicle, the mature hair, mainly consists of three differentiated cell types, the cortex, the cuticle and the medulla, and we discuss them in that order (Fig. 16.2). The cortex Cortical cells are spindle-shaped, about 100 µm long, 5-10 µm wide at their widest point and arranged in an overlapping and interdigitating fashion along the length of the fiber. The predominant proteins of hair are classified into two groups, the keratin intermediate filament (IF) proteins that form 8-10 nm filaments and the keratin associated proteins that form a matrix between the filaments. The keratin IFs consist of two families, type I and type II, each containing at least four proteins which are referred to in the older literature as the low-sulfur protein group*. During the differentiation of hair cortical cells, 8-10 keratin IF proteins are synthesized and aggregate via dimer and tetramer steps to form the keratin IF. The aggregation proceeds according to pairing rules, that is a type I chain forms a dimer with a type II chain in specific pairs. This mechanism has been extensively explored for the epidermal keratins, but only minimally for the hair keratins.3,4 In the cortical cells the keratin IFs are spaced about 10 nm apart and the spaces between them are filled by a matrix consisting of smaller keratin associated proteins (KAP) (Fig. 16.3). There are several families of KAPs, totaling of the order of 50 proteins. Typically, up to one third of their residues are either cysteine or glycine and tyrosine. The sequence of morphological changes that occur during keratin synthesis are readily observed by transmission electron microscopy. Within the cortical cells long fibrillar aggregates of IFs, termed macrofibrils, are oriented with the direction of growth and increase in size and abundance as the cells differentiate. In the early stages of macrofibril formation the * A new nomenclature has been proposed1,2 to replace the disparate terminologies and to unify the epidermal and hair keratin IF. It is used in this review. Essentially, keratin IFs are codified as either K1.n or K2.n for type I or type II IF components, respectively. The keratin associated proteins are abbreviated as KAP. Numbers attached to these symbols denote different protein families and members of these families. For example, K2.10 is a type II IF hair keratin, KAP 4 is a family of cysteine-rich proteins and KAP 6 a family of glycine/tyrosine-rich proteins.

318

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 16.2. Diagrammatic representation in cut-away view of the various cell types that make up a mature hair fiber. Note that no distinction is made between orthocortical and paracortical cell types. The exocuticle is the outermost shaded layer of the cuticle cells, which are arranged in an overlapping fashion on the surface of the fiber. Modified from Powell BC et al. In: Jolles P et al, eds. Formation and Structure of Hair. Basel: Birkhauser Verlag, 1997:59-148; reprinted with permission.

Fig. 16.3. Diagrammatic representation of the intermediate filaments (IFs, formerly called microfibrils) and matrix complex of the cortical cells of the hair. The IFs are drawn with small cores that are seen in electron micrographs of cross-sections and their center-center spacing of 10 nm is indicated.

IFs are separated from one another by a space which is later filled by a densely-staining matrix (Fig. 16.4). The interactions of the large number of KAPs during hair growth is likely to be complex and poses a daunting problem for protein chemists to unravel. At the completion of terminal differentiation the cells are virtually filled by the IF-matrix complex, although small deposits of condensed residual cytoplasm and nuclear remnants remain between the macrofibrils. Two types of cortical cells can be distinguished in the hair cortex and are termed paracortical and orthocortical cells. These two cell types are distributed bilaterally in the fine fibers of sheep (20 µm diameter or less) (Fig. 16.5) but in other species they are distributed differently, with “doughnut” and “dumbbell-shaped” patterns observed in fiber crosssections. The filaments of the paracortical cells in Figure 16.5 are arranged mainly in quasihexagonal close-packing and are clearly seen because they are “end-on”. By comparison, the IFs of orthocortical cells are closer together and form an aggregate (a macrofibril) in which the filaments become inclined to the cell axis and entwined in a rope-like fashion. Consequently, when the plane of section passes through the central region the filaments are seen

Late Events and Regulation of Keratinocyte Differentiation in Hair and Feather Follicles

Fig. 16.4. Electron micrograph of a cross-section of the developing filament-matrix complex in part of the cytoplasm of a hair cortical cell (sheep). Early aggregates of macrofibrils (M) contain regions where densely-stained matrix material clearly demarcates individual keratin IFs as seen in the keratinized state in Fig. 16.5. Other regions of the macrofibrils, indicated by arrows, are where the inclusion of matrix proteins has been incomplete and the IF-matrix structure is not visible. The dark-staining material (C) is ribosomal cytoplasm still active in protein synthesis. Bar = 100 nm. Fig. 16.5. Electron micrograph of a cross-section of a wool fiber. This picture clearly demonstrates the two distinct major types of IF-matrix packing in orthocortical (O) and paracortical (P) cells. Arrows indicate cell boundaries and between these boundaries is the band of material that is termed the δ-layer.6,21 Bar = 100 nm.

319

320

Molecular Basis of Epithelial Appendage Morphogenesis

“end-on”, whereas outwards from the core they are inclined to the section plane and appear as short lines, giving the macrofibrils the appearance of whorls in the electron microscope.5,6 The molecular basis for these packing modes is unknown, but it is likely that they are a result of differences in the distribution of the hair KAPs of the matrix. There is evidence that the ratio of IFs to matrix is higher in orthocortical cells than in paracortical cells5,7 and, as the expression of the keratin IF genes appears to be the same in all cortical cells,8,9 the altered ratio probably results from differences in particular KAPs. Indeed, the exclusive expression of the KAP4 cysteine-rich protein family in the paracortex,10 and of some glycine/ tyrosine-rich KAPs in the orthocortex,11,12 could be sufficient to account for the ultrastructural differences. The amount and composition of the matrix component of the cortex is quite variable and, in sheep, is known to be linked to nutritional status.13 In wool fibers, the higher the cystine content the greater the amount of matrix between the filaments.14,15 In sheep, supplementation with L-cysteine has a number of effects, including increases in fiber diameter and the proportion of paracortical cells.16 There is also a marked increase in the amount of the KAP4 family of cysteine-rich proteins in the paracortical cells, but no changes in other keratin families.10 The cuticle The cells that give rise to the hair cuticle undergo a morphologically and biochemically distinctive differentiation from those of the cortex. Their general shape and disposition (like tiles on a roof) on the outer surface of the fiber is diagrammatically represented in Figure 16.2. They apparently develop their outward slope and flattened shape from relative cell movements and other forces generated in the follicle during fiber growth by the innermost face of the apposed inner root sheath cuticle.17,18 Three layers are distinguishable within the flattened cuticle cell (Fig. 16.6), an outermost layer termed the exocuticle, including a narrow layer at its outer edge called the “A” layer, and an inner layer known as the endocuticle.6 The exocuticle is produced by the aggregation of protein granules which start to appear when the cuticle cells leave the follicle bulb.19,20 There are three types of granule and they congregate at the outer margin of the cuticle cell, where they consolidate to form the exocuticle late in differentiation.20 The molecular basis of this process and the conformational changes that occur in the formation of the exocuticle are not understood, including the origin of the outermost “A” layer, which is more electron-dense than the rest of the exocuticle. The electron density of the “A” layer almost certainly originates from a higher content of disulfide bonds reacting with the heavy metals used in the staining for electron microscopy because the layer also reacts more intensely with mercury compounds that bind to sulfur bonds.21 Maturation of the cuticle cells finally involves shrinkage and dehydration of the underlying flattened zone of residual cytoplasm to form the endocuticle layer. The mature cuticle cell contents are ultrastructurally homogeneous and cystine-rich.5 At least one family of proteins (KAP5), containing on average 30 mol% of both cysteine and glycine, is known to be expressed late in cuticle cell differentiation.22,23 Given the dynamics of exocuticle formation by granule accretion, it is possible that the KAP5 proteins form part of the inner exocuticle layer. The KAP5 proteins are predicted to have complex secondary structures, in which the repeating cysteine-rich and glycine-rich motifs adopt highly folded conformations.1 Unlike the cortical cells, keratin IF are not produced in great abundance in cuticle cells. A novel type I hair keratin IF, related to the keratins expressed in the cortex but with substantially different N- and C-terminal domains, is exclusively expressed in the cuticle.24 Its N-terminal domain is predicted to be more flexible than that of the cortical keratins. This cuticle type I IF gene is transcribed as the cuticle cells commence terminal differentiation, well in advance of the KAP5 genes. Several other IF proteins, normal components of

Late Events and Regulation of Keratinocyte Differentiation in Hair and Feather Follicles

321

Fig. 16.6. Electron micrograph of the cuticle of a human; hair fiber in cross-section showing seven overlapping cuticle cells. The exocuticle (exo) and its outermost “A” layer and the innermost endocuticle layer (endo) are marked in one cell. Vertical arrows show the intervening cell membrane complex (CMC), including the δ-layer. The underlying cortex is also shown (co). Bar = 0.2 µm. (Courtesy of Dr. L. Jones).

the epidermis and outer root sheath (ORS), also appear to be expressed in the cuticle,25-28 consistent with the electron microscopic observations of filamentous tufts in the early stages of cuticle differentiation.7,20 A specialized hydrophobic surface layer about 10 nm thick, the epicuticle, is present on the surface of hair fibers. The origin and chemistry of this surface layer have been obscure for many years5,29 but significant advances have been made recently. The principal lipid responsible for the hydrophobicity is the C21 saturated fatty acid 18-methyl-eicosanoic acid,30 which is covalently associated with a surface protein by a thioester linkage with exposed cysteine residues.31,32 This surface layer (see below) is produced in the environment of the “cell membrane complex” (CMC), the intercellular lamina (see Figs. 16.5 and 16.6) that develops between the plasma membranes of the cortical cells as well as the cuticle cells5 and was originally called the δ-layer.6,21 The medulla The medulla (Fig. 16.1) is frequently present in animal hairs but does not occur in fine wool fibers. With its entrapped air, the medulla provides insulating capacity to the hairs of the pelage. In some species it is a continuous structure and can represent more than 15% of fiber weight,33 but in others, for example humans, it can be discontinuous. The medulla is centrally placed within the cortex and is made up of solidified cells that vary in their number and arrangement according to species. The cells often form a column in a regular girderlike arrangement, wedged between “trabeculae” which are projections from adjacent and deformed cortical cells. Ryder34 has classified the various arrangements of medulla cells in the hair of different species.

322

Molecular Basis of Epithelial Appendage Morphogenesis

Medulla cells arise from a differentiation program that is distinct from that of the cortex. As medullary cells leave the bulb region, moving upwards from the top of the papilla, they begin to differentiate and synthesize the protein trichohyalin. Trichohyalin deposits in the cytoplasm as granules but does not completely fill the differentiated cells so that on maturation and dehydration the cells shrink and become wedged between the cortical “trabeculae” with air spaces between them. The medullary trichohyalin granules are morphologically and histochemically very similar to the trichohyalin granules in the IRS, but suffer a different fate.35 In mature medulla cells the trichohyalin is a mass of fused protein granules, whereas in the IRS the trichohyalin granules disperse between filaments during the later stages of differentiation (see below). Trichohyalin contains citrulline residues and is crosslinked by isopeptide bonds36,37 instead of disulfide bonds, typical of the surrounding hair cortex.38,39 The inner and outer root sheaths of the hair follicle The IRS and ORS layers of the follicle are intimately associated with the growth and maintenance of hair, although they do not synthesize hair keratins. The differentiation pathway in IRS cells produces a hardened layer but the participating proteins are markedly different from that of hair. The ORS is a multilayered cylindrical structure that envelopes the IRS and through which there is exchange of metabolites between the growing hair and the surrounding dermis and capillary system. The three cellular layers of the IRS, the IRS cuticle, Huxley layer and outermost Henle layer, are believed to originate from germinative cells near the base of the dermal papilla. Desmosomes are abundant between the cells of the IRS.5,17,39 The IRS cells move up the follicle inside the surrounding ORS, physically supporting the growing fiber, but above the sebaceous gland the IRS cells dissociate and slough into the pilary canal, releasing the emerging hair. The earliest sign of differentiation in the IRS is the appearance of trichohyalin granules in association with filaments 8-10 nm in diameter, but these filaments are not the same as those that are found in the hair cortex and discussed elsewhere. When the IRS cells have completely differentiated no granules are visible, only the 8-10 nm diameter filaments oriented in the direction of fiber growth.40 It was once thought that the granules gave rise to the filaments, but it is now accepted that they form a matrix between the filaments.39,41-43 Although the filaments are not of the hair type, epidermal keratin proteins including K1.16, K2.1, K2.6 and possibly K1.10 have been detected in IRS cells.26,28 K2.1 and K1.10, and K2.6 and K1.16, are typical partners in filament formation and therefore may constitute the filaments that appear later in IRS differentiation, but this is not proven. The trichohyalin gene has been cloned from sheep and human.41,43 The sheep protein contains 1549 residues43 and the sequence of human trichohyalin is similar.41 The obligatory 315 residue α-helical domain of heptad repeats characteristic of IFs is absent and hence trichohyalin is not a precursor of an IF but is an IF-associated protein.43 The sequence has an abundance of glutamic acid, arginine, glutamine and lysine residues, the latter three being substrates for the enzymes peptidylarginine deiminase (PAD: arginyl residues) and transglutaminase (glutaminyl and lysyl residues). They are arranged in 28 amino acid repeats in the center of the molecule and in 23 amino acid repeats at the C-terminal end. At the N-terminal end of the protein there are two regions of sequence of about 28 amino acid residues each that have a helix-turn-helix conformation homologous with that found in calmodulin and related proteins which bind calcium strongly.43 These sequences are called E-F hands and each E-F hand is capable of binding a calcium ion. Since the trichohyalin molecule has two E-F hands, it can therefore potentially bind two calcium ions and this has been demonstrated experimentally.41

Late Events and Regulation of Keratinocyte Differentiation in Hair and Feather Follicles

323

The differentiation of the IRS cells involves two major chemical changes that are crucial for the formation of the filament-matrix complex and the conversion of the cellular protein to an insoluble and hardened state. The PAD enzyme acts on the basic arginine residues of trichohyalin, converting them to neutral citrulline residues.44 This reaction is believed to cause a radical decrease in ionic side chain interactions in trichohyalin and a conformational change that leads to its dispersion between the filaments as a matrix protein.40,42,43 The strategic co-location of PAD in relation to trichohyalin has been reported for the IRS and, as expected, the relationship holds for the cells of the medulla as well.45 The second chemical change is that of isopeptide crosslinking between glutaminyl and lysyl residues, which accounts for the final insolubility of the hardened cell contents. The transglutaminase enzyme responsible for this event was first detected in hair follicles by Harding36 and shown to be present in the IRS cells.46 The ORS is unlike other concentric cell layers of the follicle in that it has its own autonomous cell population. The ORS is continuous with the epidermis, so it is not surprising that epidermal and not hair type IFs are expressed. The basal keratin pair in the epidermis, K2.5 and K1.14, is also abundant in the ORS. The pattern of K1.14 expression above the sebaceous gland is similar to that found in the epidermis, where it is expressed in the mitotically active basal cells.47 It is curious, then, that in the part of the ORS that extends down from the sebaceous glands to the base of the follicle, K1.14 is expressed in the inner, suprabasal ORS cells. K1.17 is also found in the ORS28,48 and K1.18, K1.19 and K2.8 are present in low abundance in the lower ORS in a mosaic of expressing and nonexpressing cells.26 The innermost ORS layer, next to the Henle layer, has a CMC with a central δ-layer thinner than that between other follicle cells.39 Orwin17 named this layer of ORS cells the companion layer because it appeared to accompany the IRS cells in their upward progression.

Growth and Late Biochemical Events in the Hair Follicle The normal rates of growth of hair fibers are about 300 µm per day and are determined by the rate of supply of cells from the follicle bulb and their distribution between the various layers of the fiber and IRS. It has been estimated that no more than 20% of the bulb cells differentiate into the fiber cortex, the rest forming the IRS.49 The mechanisms determining cell fate in the follicle bulb are not known, but cell-cell interactions are undoubtedly important. For example, the mouse gene for Notch, a transmembrane protein important in the process of cell fate selection,50 is expressed in all follicle bulb cells except for those apposed to the dermal papilla.51 This location accords with a model in which neighboring follicle bulb cells interact via Notch and enter their respective differentiation pathways to produce the different follicle cell types. In relation to this hypothesis, it is notable that adjoining cells in the follicle bulb have numerous gap junctions and desmosomes in their membranes17 and some cells appear to be compartmentalized into groups by boundaries that restrict the passage of molecules from one group to another.52 Dynamic changes in cell boundaries and intercellular communication might be crucial in determining the fate that cells take on leaving the follicle bulb. As they move upwards from the follicle bulb the cortical cells begin to synthesize keratin proteins, increasing to a maximal rate in the lower third of the follicle. The availability of adequate supplies of essential amino acids is vital for normal hair growth and this is dramatically apparent in sheep because there is a very large population of follicles competing for amino acid substrates.53 A deficiency in the supply of sulfur amino acids results in lower cell division rates in the follicle germinative cells, a reduced rate of hair growth and a change in the pattern of keratin synthesis.10,16 The terminal stages of cortical cell differentiation include removal of the cytoplasm, cell organelle destruction and finally dehydration of the cell. The degradative events are

324

Molecular Basis of Epithelial Appendage Morphogenesis

virtually completed at the level of hardening of the hair fiber, but very little is known about the processes involved. It is uncertain as to whether DNA is completely removed from the nuclear remnant.54-56 The establishment of disulfide cross-links between the numerous protein components, all of which contain cysteine residues, is a major event in the terminal differentiation of hair keratinocytes, which is not understood. The hair proteins are maintained in the sulfhydryl state during the period of protein synthesis,57 and it has been proposed that the reducing potential required to maintain this state is provided by NADPH.58 At the level of fiber hardening in the follicle (see Fig. 16.1) a rapid oxidative step extending over a length of several cortical cells produces an almost completely disulfide cross-linked keratin complex in the fiber. Sulfhydryl concentration in the wool follicle reduces from about 1100 micromoles/g (calculated for wool of 3.5% sulfur content) before oxidation to 12-30 micromoles/g after oxidation.13 The mechanism of oxidation is uncertain, although it appears that copper ions or a copper-containing enzyme are responsible for catalyzing the reaction. A mutation in a copper transporter protein has recently been shown to be responsible for Menkes syndrome,59 a human condition manifested by kinked hair in which the sulfhydryl content of the hair is only partly oxidized to disulfide bonds.13 A sulfhydryl oxidase that might be involved in this process has been isolated from rat skin by Takamori et al60 and evidence suggests that this enzyme is active in hair follicles, but no involvement of copper was mentioned. During the terminal stages of differentiation an intercellular CMC appears between cortical and cuticle cells and at the junctions between cortex and cuticle.39,61-63 A CMC also forms an adherent layer at the junction of the hair cuticle surface and the IRS cuticle. The separation of the fiber from the IRS cuticle and the disappearance of the IRS without any apparent effect on the hair itself are intriguing processes. Since the cells of the developing fiber and those of the IRS are well advanced towards terminal differentiation at this stage it appears that the sebaceous gland is a source of proteases that carry out the selective digestion releasing the fiber surface from the IRS cells.64 Compared to the cell membranes in the lower regions of the follicle that contain various phospholipids typical of living cells, the CMC of mature wool fibers has little phospholipid and mainly consists of free fatty acids and the sterols cholesterol and desmosterol.65 There is good evidence that the CMC proteins are cystine-rich7,29 and are also crosslinked by isopeptide bonds.66 Presumably there must be physical attachments of hair keratin IF to the CMC, but they have not been described. Glycoproteins have also been reported in the CMC.67

The Genes for Structural Proteins and Their Expression in the Cortex and Cuticle During Hair Growth Knowledge of the complexity of hair keratin proteins as large gene families has increased enormously in recent years through molecular studies, and many genes for IFs and KAPs have been isolated. The structure of the hair keratin IF genes is similar to the epidermal IF genes, pointing to common evolutionary ancestors. The type I IF genes contain 6 introns and are 4-5 kb in size whereas the type II IF genes contain 8 introns and are 7-9 kb in size.8,9,68,69 Genetic and chromosomal mapping data indicate that various linkages exist between different keratin gene families. Hair and epidermal keratin IF genes appear to be linked but the type I genes are located on a separate chromosome from the type II genes.70-72 There are some tantalizing exceptions to this general arrangement, however. The K1.18 gene appears to map to the type II locus in humans,73 and various epidermal keratin probes show hybridization to other chromosomal locations,72,74,75 possibly representing bona fide “orphan” keratin genes or pseudogenes.

Late Events and Regulation of Keratinocyte Differentiation in Hair and Feather Follicles

325

Information on type II IF gene organization is more advanced than for the type I genes. A 630 kb YAC clone has been isolated containing eight human epidermal type II IF genes, K2.1–K2.8 and several related genes,73,76 and cosmid data indicates close linkage of three hair and three hair-related genes within a 100 kb contig in the sheep genome.9 Predictions based on these data suggest that the type II locus might span 500-600 kb and encompass 20-30 genes. Individual clusters of hair68,77,78 and epidermal keratin type I IF genes74,79,80 have also been described. Linkage between specific hair and epidermal keratin IF genes has yet to be demonstrated, however. The hair KAP genes are more numerous than the IF genes; there are more than ten families, and in general the genes within each family appear to be tightly clustered, frequently within a few kb of each other (for review see ref. 1). Recent studies indicate that in sheep the KAP1 gene family is linked to the type I IF locus.81 Other mapping data (Powell et al unpublished) suggests that the KAP1 and KAP2 families may be linked, raising the possibility of a large cluster of type I IF and hair KAP gene families. Given the probable common evolutionary origin of the chromosome segments that include the two keratin IF loci,82,83 it is possible that hair KAP genes may also be linked to the type II IF gene locus. However, some hair KAP genes are known to map to different chromosomes; the cuticle KAP5 genes map to human chromosome 1584 and the KAP6 gene family and the KAP8 gene, encoding glycine-tyrosine-rich hair proteins, map to sheep chromosome 1.81,85 Immunochemical studies of keratin formation have been carried out, in all cases with antibodies that were probably pan-specific.26,86-90 More precise information on the expression of hair keratin genes has been obtained in the last few years using in situ hybridization with specific cRNA probes. These studies have indicated that the genes of the three major classes of proteins, the keratin IF, the glycine/tyrosine-rich and the cysteine-rich proteins are expressed sequentially during hair growth in that order. The hair keratin IF proteins (Fig. 16.7) are the first differentiation-specific proteins produced,8,24,68,91-94 type II IF genes, for example, being expressed sequentially in cortical cell differentiation in separate but closely-timed stages8,94 and presumably in parallel with their type I partners. As mentioned earlier, a new type I IF gene is exclusively expressed in the cuticle.24 The mRNA first appears in cuticle cells situated around the periphery of the upper part of the follicle bulb. This type I IF gene is therefore one of the first differentiationspecific genes expressed in cuticle cells and in its timing of expression is comparable to K2.12, a type II IF keratin gene of the cortex.1,94 In the last few years some interesting features of the KAP genes that encode the proteins of the matrix have emerged. There are at least 10 families of KAP genes, ranging in size from 0.6 to 1.5 kb. A striking feature of their structure is the absence of introns, a rare occurrence in vertebrate genes, and the hair KAP genes would constitute one of the largest groups of intron-less genes known. In addition to the KAP genes that produce the matrix proteins of the cortical cells, a different KAP family (KAP5) is expressed in the human hair cuticle. The complexity of KAP gene expression in the hair cortical cells is probably an order of magnitude greater than that of the keratin IF genes because of the numbers of genes involved. Currently, there are published expression data for the cortical KAP4, 6, 9 and KAP11 genes and the cuticle KAP5 genes.10,12,22,23,95-97 The patterns of expression of the KAP genes have been most comprehensively studied in the wool follicle. Following the onset of expression of the hair keratin IF genes in the cells of the mid bulb region, the KAP genes are sequentially activated in different patterns in the lower to mid-follicle shaft (Fig. 16.8). The genes encoding the glycine/tyrosine-rich KAPs (KAP6 family, KAP7 and KAP8 proteins) are expressed first, and their proteins could be the first proteins of the matrix to interact

326

Molecular Basis of Epithelial Appendage Morphogenesis Fig. 16.7. In situ hybridization of a cRNA probe localizing the expression of an IF gene in the cortex of a wool fiber. The expression begins just above the bulb (B, and arrow) and extends for the lower third of the follicle length. Bar = 54 µm.

with the nascent filaments, perhaps helping to establish the orderly spacing of the filaments98 prior to the influx of the cysteine-rich proteins that are synthesized later. In sheep Merino follicles, that show a marked bilateral segmentation of the cortex into orthocortical and paracortical cells, the expression of the KAP 6, 7 and 8 genes encoding glycine- and tyrosine-rich proteins is restricted to cells of the orthocortical lineage. The KAP4 family encoding cysteine-rich proteins is expressed slightly later and appears to be restricted to paracortical cells.10 Two other KAP gene families encoding different cysteinerich proteins, so far only described in the mouse, are expressed in all cortical cells of the lower shaft.96,97 In the cuticle, the KAP5 family encodes proteins which are cysteine-rich and also glycine-rich, and which are expressed late in hair cuticle differentiation.22,23 A new cuticle-specific gene family, KAP10, has been discovered very recently and is also expressed late in cuticle cell differentiation (Powell et al, in preparation). The encoded protein is cysteine-rich but has a completely different composition to the cuticle KAP5 proteins, with virtually no glycine. These proteins appear at similar times in differentiation and it will be interesting to find out where they are incorporated into the ultrastructure of the cuticle cells.

Late Events and Regulation of Keratinocyte Differentiation in Hair and Feather Follicles

Cuticle Paracortex

KAP5 & KAP10 KAP4

Cortex

KAP1, 2, & 3

Orthocortex

KAP6, 7 & 8

Cortex

327

Keratin type I & type II IF

Fig. 16.8. Cartoon of the hair follicle summarizing the regional and temporal differences in keratin gene expression. Expression is detected by experiments using specific cRNA probes. The data for this cartoon have been derived from observations on both wool follicles and hair follicles of other species. Modified from Powell BC et al in: Jolles P et al, eds. Formation and Structure of Hair. Basel: Birkhauser Verlag, 1997:59-148; reprinted with permission.

Transcriptional Control of Gene Expression The hair keratin IF and KAP genes are expressed in a cell type-specific and temporallycontrolled manner during follicle differentiation and the molecular mechanisms which control this process are likely to be complex. One factor which appears to figure prominently in the control of hair keratin gene expression and which, surprisingly, is also important in the early stages of follicle development, is lymphoid enhancer factor 1.99 A comparison of fifteen hair keratin gene promoters reported by Powell et al100 revealed several highly-conserved DNA sequences grouped with particular gene families. One of these is identical to the consensus binding site CTTTGA/TA/T for lymphoid enhancer factor 1 (Lef1).99 This site is also present within the proximal promoters of hair KAP genes. Lef1 contains a DNA-binding domain related to that of the high mobility group proteins, and binding of Lef1 has been shown to cause a bend in the DNA helix of the T cell receptor α and HIV-1 enhancers, facilitating protein-protein interactions.101-103 Possibly, Lef1 may perform a similar function in regulating the hair keratin gene promoters, promoting cooperative binding of factors required for high level gene expression. Its ability to promote cooperative transcription factor interactions poses a question as to what factors it might interact with in regulating hair keratin gene expression. In the T cell receptor α and HIV-1 enhancers, the Lef1 site is positioned between binding sites for strong, inducible activators (NF-κB for HIV-1 and CREB/ATF for TCR-α) and the factor Ets-1. The interaction of these sites appears to be essential for enhancer function.102,103 In an analogous situation, in the K2.10 IF promoter the Lef1 site is flanked by potential Ets-1 and Sp1 binding sites on one

328

Molecular Basis of Epithelial Appendage Morphogenesis

side and NF1 sites on the other (Fig. 16.9). Binding of factors to these sites and their interaction via Lef1 may prove to be important in regulating this hair keratin gene. Significantly, mutagenesis of the Lef1 binding site in the hair K2.10 promoter severely reduced the level of expression in transgenic mice, consistent with a vital role for Lef1 in enhancing hair keratin gene expression (Keough, Powell and Rogers, unpublished data). In addition to its role in regulating hair keratin gene expression, Lef1 performs other important functions in the hair follicle. In Lef1 knockout mice, follicle development is arrested not long after initiation, indicating an essential role for Lef1 before its involvement with hair keratin genes.104 Lef1 is also expressed in different patterns in embryonic vibrissae and postnatal pelage follicles.99 In vibrissae follicles, Lef1 and hair keratin IF genes appear to be coexpressed, whereas in pelage follicles Lef1 expression is detectable in bulb cells apparently prior to that of the hair keratin genes. Lef1 is also expressed in the dermal papilla, which does not express hair keratin genes. At this stage it is not known whether Lef1 is also expressed later in hair follicle differentiation in the mid hair shaft but, given the presence of Lef1 sites in hair KAP genes that are not activated until later, Lef1 is expected to be present throughout hair cortical keratinocyte differentiation. To investigate promoter function in hair keratin genes it is currently necessary to use transgenic mice because of the lack of a cell culture system. Two such studies have shown that faithful hair-specific expression of a heterologous reporter gene is maintained when either a 400 bp region of the hair keratin gene K2.10 promoter2 or a 600 bp region for a matrix gene promoter was used.96 Since hair follicles are derived from the epidermis, some of the features of the control of epidermal keratin genes are likely to be involved in hair keratin gene expression. Nevertheless, there must also be distinct gene controls operating in the hair follicle that direct expression in separate regions. For example, epidermal keratin genes are expressed in the outer and inner root sheaths, whereas hair keratin genes are expressed in the fiber cortex. Similarly, there is expression of the two types of keratin genes in separate compartments in the filiform papillae of the tongue105 and in nail.106 The study of regulatory elements in epidermal keratin genes has advanced more rapidly because of the availability of appropriate cell lines. Many factors have been shown to influence epidermal differentiation and keratinization, including calcium ions, retinoids, vitamin D3 and growth factors such as EGF, TGF-α and TGF-β.107-111 Epidermal keratin genes that are influenced by these signals have been analyzed for regulatory elements in attempts to identify the transcription factors involved, as well as factors which dictate keratinocyte specificity. The transcription factor AP2, which is expressed in many tissues including the epidermis,112 seems to regulate the activity of several keratin IF genes. K1.14 and K2.5, which are coordinately expressed in the basal cells of the epidermis and the follicle ORS, both bind AP2.113,114 Leask et al115 identified two regions of the human K1.14 gene promoter which act cooperatively to promote keratinocytespecific expression in cultured cells and transgenic mice. A distal element, as yet uncharacterized, is located between -1700 and -2100 bp upstream of the transcription start site and a proximal element which binds AP2 is located between -160 and -270 bp. Although the AP2 binding site is important for K1.14 expression, high level expression requires the additional distal enhancer.114 AP2 also binds to the promoter of the K2.5 gene, which is coordinately expressed with K1.14.113,114 Given that AP2 binds the K1 (K2.1) promoter, which is expressed in the suprabasal layer of the epidermis and in the follicle IRS, this transcription factor may be employed throughout much of the epidermis and its derived structures.114 Other common transcription factor binding sites found in epidermal keratin genes include those for AP1, Sp1, and NF1 proteins, as well as steroid/thyroid hormone nuclear receptors.116-119 It is increasingly apparent that keratinocyte-specific transcriptional

Fig. 16.9. Putative regulatory motifs in the promoter region of a hair keratin IF type II gene (sheep K2.10). This sequence confers folliclespecific expression of a reporter gene in transgenic mice (Keough, Powell and Rogers, unpublished data). The location of the transcription start site is indicated at +1 bp. In addition to the CAAT and TATA motifs, various potential binding sites for AP2, Sp1, Ets-1, NF1 and Lef1 are also shown.

Late Events and Regulation of Keratinocyte Differentiation in Hair and Feather Follicles 329

330

Molecular Basis of Epithelial Appendage Morphogenesis

control arises by specific association of two or more factors interacting with clusters of binding sites in the promoters or enhancers. In the human K2.5 gene, for example, a promoter sequence with keratinocyte-specific activity contains binding sites for Sp1, AP2 and a number of other factors within approximately 100 bp of the transcription start site.113 Casatorres et al120 also described an epithelial-specific enhancer in the K2.5 gene which seems to function via an AP1 element. Since AP1 expression is widespread, it was suggested that specificity for keratinocytes is conferred by additional sequences and possibly by other protein interactions within this enhancer. Another example of the interplay of transcription factors comes from the human K2.1 gene.110 K2.1 expression in the epidermis is initiated in differentiating suprabasal cells and is regulated by calcium levels, vitamin D3 and retinoic acid. The calcium response element, located 3' of the K2.1 gene,121 contains an AP1 binding site and the adjacent sequence contains a hormone responsive element through which activated receptors of vitamin D3 and retinoic acid modulate the calcium response.110 The gene for involucrin, a major structural component of the epidermal cornified envelope, is expressed in the suprabasal layer of the epidermis and is also expressed at high levels in the follicle IRS and to a lesser extent in the ORS and hair cortex.122 The involucrin promoter is activated by AP1,123,124 and a factor which binds an AP2-like motif has also been implicated in differentiation-dependent expression.125 Welter et al126 demonstrated that the activity of the involucrin promoter could be suppressed by various POU domain proteins. Transcription of the keratin IF genes K1.14 and K2.5 is also apparently suppressed by POU proteins.127 In both studies, suppression did not seem to involve direct binding of the POU factors to the promoters and they presumably exert their effects by regulating other transcription factors, either via direct interaction or indirectly by regulating their expression. Another gene expressed in the IRS and medulla of the hair follicle and in the late stages of epidermal differentiation encodes the enzyme transglutaminase III which crosslinks trichohyalin. Dissection of the proximal promoter of this gene has indicated a requirement for adjacent Sp1 and Ets-1 binding sites for expression in epithelial cells.128 In this context it is interesting to note that there are potential Sp1 and Ets-1 binding sites in the minimal hair K2.10 promoter (Fig. 16.9) and that Ets-1 is expressed in the hair follicle.129 These factors may therefore control the expression of genes in several different compartments of the hair follicle. Potential binding sites for AP1 and AP2 are also present in many hair keratin gene promoters8,9,12,100 and the finding of Fos, one of the components of the AP1 complex, in the hair follicle cortex and the IRS130 supports the likely role of this factor in regulating hair genes. In a search for novel transcription factors, Andersen et al131 identified two factors, Skn-1a and Skn-1i, expressed by alternative splicing. These proteins are closely related to Oct-11 and contain the bipartite POU domain DNA binding motif common to this family.132 Skn1a, the active form of the protein, appears to enhance transcription from the K1.10 promoter.131 In situ hybridization analyses of Skn-1a/i expression showed that they are expressed in the suprabasal cells of the epidermis and in differentiating cortical cells of adult hair follicles, suggesting that they might be important in the late stages of the keratinization of hair. The evidence to the present time has not revealed any unique transcription factors controlling the expression of hair keratin genes, although the data must be considered preliminary at this stage since the study of hair keratin IF and KAP gene regulation is just commencing. Given the redundancy of transcription control in other systems, where finetuning of gene expression can be brought about by subtle changes in factor interaction and binding affinity for a promoter, it is possible that there might not be factors that are unique to the hair follicle.

Late Events and Regulation of Keratinocyte Differentiation in Hair and Feather Follicles

331

Comparison of Hair Keratinization with Avian Keratin Formation in the Feather Follicle The feather follicle is a useful system for studying morphogenesis and terminal differentiation of keratinizing appendages, as discussed in Part II. In comparison with hair formation, in which diverse families of proteins are expressed in distinct cell streams during differentiation, the feather follicle is a simpler system. Feathers are derived from a family of perhaps twenty avian β-keratins that are similar to one another in amino acid sequence. The avian β-keratins are smaller molecules (10 kDa) and differ completely in composition, sequence and structure from the proteins of hair.5,133 Their genes apparently evolved by gene duplication.134,135 The avian β-keratins include those of the scales, beaks and claws as well as feathers. The keratins of scale, claw and beak contain the feather keratin sequence but with repeated inserts of short peptide sequences (9-13 amino acid residues) rich in glycine and tyrosine.136,137 In the evolutionary development of feathers from scales the additional peptide sequences appear to have been deleted.136,137 The 1 kb-long feather keratin genes are clustered and tandemly organized with a center to center separation of 3.3 kb, and transcription of at least four of the approximately twenty genes has been demonstrated.135,138 The expression of feather keratin increases rapidly after the formation of the feather bud at about 12 days of embryonic development and reaches a maximal rate 2-3 days later.139 In addition, autoradiographic electron microscopy showed that the mRNA-ribosome machinery is attached to the filaments, from which it was concluded that β-keratin chains aggregate quickly until the dimensions characteristic of the filaments (4 nm diameter) are reached.139 In itself, the determination of filament size is another interesting phenomenon for which the structural explanation is not yet apparent. Compared to the hair follicle, even less is known of the molecular controls that regulate avian β-keratin synthesis after the early cylindrical follicle has been formed and before the sheath breaks to produce the mature feather. For example, the regional distribution of expression of the different genes is not known nor what elements control their transcription or whether any cellular factors are involved in the aggregation of the β-keratin chains to form filaments. Comparative analyses revealed several regions of high sequence conservation in the 5' and 3' noncoding regions and in the intron of the feather keratin genes, one 18 bp segment of the 3' noncoding region, for example, being perfectly conserved amongst five genes (for review see ref. 140). Experiments in the Xenopus oocyte transcription system indicated that a negative control element resides in the 5' intron of feather genes,141 perhaps located in one of the conserved regions. A clue to possible cellular factors active in the formation of feather keratin filaments lies in the finding of another protein present in the keratinizing feather. This 14 kDa protein (possibly a protein family) was originally called “fast protein” (Fp) because of its high electrophoretic mobility, a result of its histidine and arginine content.142 It is found in embryonic feathers and scales but is absent from the adult structures.140 The 5' noncoding region of the Fp gene has regions that are homologous with those of the feather keratin genes, suggesting that they might share some control mechanisms.140 A more recent study of Fp (where it is referred to as histidine-rich protein B or HRP-B) has defined its cellular location during embryonic development of both feather and scale by immunohistochemistry using a polyclonal antibody.143 HRP-B is found in the barbs, barbules and medulla cells at the same time as feather keratin. It is not present in nonfollicular epidermis. In scutate scales, it is only found in the very early embryonic layers and not in the later stages of keratinization. HRP-B therefore appears to have a strong association with the synthesis of feather keratin, but whatever that molecular role is, the protein disappears by the time the adult feather is formed.

332

Molecular Basis of Epithelial Appendage Morphogenesis

Concluding Remarks It is clear from this review of the field that there is still much to be revealed about the molecular events that occur during keratinocyte differentiation and which lead to the synthesis of specific families of keratin filament proteins together with their particular associated proteins. Different protein sets are produced in the many kinds of epithelial appendages. Thus, in the cortex of the hair follicle the specific hair keratin filaments are associated with special matrix proteins, whereas in the inner root sheath the filaments are associated with trichohyalin as the matrix. In the epidermis, a different set of keratin filaments are produced and associate with filaggrin. In the feather follicle and chicken scale it appears that the β-keratin proteins are sufficient to form the hardened structures by themselves and that the filament-matrix arrangements found in mammalian tissues are not necessary. We have little detailed knowledge of the genomic mechanisms responsible for the selection of particular keratin genes for expression. A major challenge for molecular biologists is to understand how the keratin genes are regulated to produce the orderly sequence of synthesis of keratin proteins that accompanies the formation of the appendages and their maintenance, as exemplified in the hair follicle.

Acknowledgments The research work of the authors reproduced in this review has been supported by grants from the Australian Wool Research and Promotion Organization/International Wool Secretariat and from the Australian Research Council.

References 1. Powell BC, Rogers GE. The role of keratin proteins and their genes in the growth, structure and properties of hair. In: Jolles P, Zahn H, Hocker H, eds. Formation and Structure of Hair. Basel: Birkhauser Verlag, 1997:59-148. 2. Rogers GE, Powell BC. Organization and expression of hair follicle genes. J Invest Dermatol 1993; 101:50S-55S. 3. Coulombe PA, Fuchs E. Elucidating the early stages of keratin filament assembly. J Cell Biol 1990; 111:153-169. 4. Herrling J, Sparrow LG. Interactions of intermediate filament proteins from wool. Int J Biol Macromol 1991; 13:115-119. 5. Fraser RDB, MacRae TP, Rogers GE. Keratins. Their Composition, Structure and Biosynthesis. Springfield, Illinois: Charles C. Thomas, 1972. 6. Rogers GE. Electron microscopy of wool. J Ultrastruct Res 1959; 2:309-330. 7. Marshall RC, Orwin DFG, Gillespie JM. Structure and biochemistry of mammalian hard keratin. Electron Microsc Rev 1991; 4:47-83. 8. Powell B, Crocker LA, Rogers GE. Hair follicle differentiation: Expression, structure and evolutionary conservation of the hair type II keratin intermediate filament gene family. Development 1992; 114:417-434. 9. Powell BC, Beltrame JS. Characterization of a hair (wool) keratin intermediate filament gene domain. J Invest Dermatol 1994; 102:171-177. 10. Fratini A, Powell BC, Hynd PI et al. Dietary cysteine regulates the levels of mRNAs encoding a family of cysteine-rich proteins of hair. J Invest Dermatol 1994; 102:178-185. 11. Hewish DR, French PW. Monoclonal antibodies to a subfraction of merino wool hightyrosine proteins. Aust J Biol Sci 1986; 39:341-351. 12. Fratini A, Powell BC, Rogers GE. Sequence, expression and evolutionary conservation of a gene encoding a glycine/tyrosine-rich keratin-associated protein of hair. J Biol Chem 1993; 268:4511-4518. 13. Gillespie JM. The structural proteins of hair: isolation, characterization and regulation of biosynthesis. In: Goldsmith LA, ed. Physiology, Biochemistry and Molecular Biology of the Skin. Vol I. Second edition. Oxford: Oxford Uni Press, 1991:625-659.

Late Events and Regulation of Keratinocyte Differentiation in Hair and Feather Follicles

333

14. Gillespie JM, Reis PJ. The dietary-regulated biosynthesis of high-sulfur wool proteins. Biochem J 1966; 98:669-677. 15. Kaplin IJ, Whiteley KJ. An electron microscope study of fibril:matrix arrangements in high and low crimp wool fibers. Aust J Biol Sci 1978; 31:231-240. 16. Hynd PI. Factors influencing cellular events in the wool follicle. In: Rogers GE, Reis PJ, Ward KA, Marshall RC, eds. The Biology of Wool and Hair. London: Chapman and Hall, 1989:169-184. 17. Orwin DFG. The cytology and cytochemistry of the wool follicle. Int Rev Cytol 1979; 60:331-374. 18. Straile WE. Root sheath-dermal papilla relationships in the control of hair growth. In: Lyne AG, Short BF, eds. Biology of Skin and Hair Growth. Sydney: Angus and Robertson, 1965:35-37. 19. Swift JA. The histology of keratin fibers. In: Asquith RA, ed. Chemistry of Natural Protein Fibers. New York: Plenum Press, 1977:81-146. 20. Woods JL, Orwin DFG. Studies on the surface layers of the wool fiber cuticle. In: Parry DAD, Creamer LK, eds. Fibrous Proteins: Scientific, Medical and Industrial Aspects. Vol 2. London: Academic Press, 1980:141-150. 21. Rogers GE. Electron microscope studies of hair and wool. Ann NY Acad Sci 1959; 83:378-399. 22. Jenkins BJ, Powell BC. Differential expression of genes encoding a cysteine-rich keratin family in the hair cuticle. J Invest Dermatol 1994; 103:310-317. 23. MacKinnon PJ, Powell BC, Rogers GE. Structure and expression of genes for a class of cysteine-rich proteins of the cuticle layers of differentiating wool and hair follicles. J Cell Biol 1990; 111:2587-2600. 24. Winter H, Siry P, Tobiasch E et al. Sequence and expression of murine type I hair keratins mHa2 and mHa3. Exp Cell Res 1994; 212:190-200. 25. Heid HW, Moll I, Franke WW. Patterns of expression of trichocytic and epithelial cytokeratins in mammalian tissues. II. Concomittant and mutually exclusive synthesis of trichocytic and epithelial cytokeratins in diverse human and bovine tissues (hair follicle, nail bed and matrix, lingual papilla and thymic reticulum). Differentiation 1988; 37:215-230. 26. Heid HW, Moll I, Franke WW. Patterns of expression of trichocytic and epithelial cytokeratins in mammalian tissues. I. Human and bovine hair follicles. Differentiation 1988; 37:137-157. 27. Bertolino AP, Checkla DM, Heitner S et al. Differential expression of type I hair keratins. J Invest Dermatol 1990; 94:297-303. 28. Stark H-J, Breitkreutz D, Limat A et al. Keratins 1 and 10 or homologues as regular constituents of inner root sheath and cuticle cells of the human hair follicle. Eur J Cell Biol 1990; 53:359-372. 29. Leeder JD. The cell membrane complex and its influence on the properties of the wool fiber. Wool Science Rev 1986; 63:3-35. 30. Evans DJ, Leeder JD, Rippon JA et al. Separation and analysis of surface lipids of the wool fiber. Proceedings of the 7th International Wool Textile Research Conference, Tokyo 1985; 1:181-193. 31. Negri AP, Cornell HJ, Rivett DE. The nature of covalently bound fatty acids in wool fibers. Aust J Agric Res 1991; 42:1285-1292. 32. Negri AP, Cornell HJ, Rivett DE. A model for the surface of keratin fibers. Textile Res J 1993; 63:109-115. 33. Bradbury JH, Leeder JD. Keratin fibers. IV. Structure of the cuticle. Aust J Biol Sci 1970; 23:843-854. 34. Ryder ML. A survey of the gross structural feature of protein fibers. In: Hearle JWS, Peters RH, eds. Fiber Structure. London: Butterworths and The Textile Institute, 1963:534-566. 35. Rogers GE. The localization and significance of arginine and citrulline in proteins of the hair follicle. J Histochem Cytochem 1963; 11:700-705. 36. Harding HWJ, Rogers GE. ε-(γ-glutamyl) lysine cross-linkage in citrulline protein fractions from hair. Biochemistry 1971; 10:624-630.

334

Molecular Basis of Epithelial Appendage Morphogenesis

37. Harding HWJ, Rogers GE. The occurrence of the ε-(γ-glutamyl) lysine crosslink in the medulla of hair and quill. Biochim Biophys Acta 1972; 257:37-39. 38. Rogers GE, Harding HWJ, Llewellyn-Smith IJ. The origin of citrulline-containing proteins in the hair follicle and the chemical nature of trichohyalin, an intracellular precursor. Biochim Biophys Acta 1977; 495:159-175. 39. Rogers GE. Isolation and properties of inner root sheath cells of hair follicles. Exp Cell Res 1964; 33:264-276. 40. Rogers GE, Fietz MJ, Fratini A. Trichohyalin and matrix proteins. Ann N Y Acad Sci 1991; 642:64-81. 41. Lee S-C, Kim I-G, Marekov LN et al. The structure of human trichohyalin. J Biol Chem 1993; 268:12164-12176. 42. Rothnagel JA, Rogers GE. Trichohyalin, an intermediate filament-associated protein of the hair follicle. J Cell Biol 1986; 102:1419-1429. 43. Fietz MJ, McLaughlan CJ, Campbell MT et al. Analysis of the sheep trichohyalin gene: Potential structural and calcium-binding roles of trichohyalin in the hair follicle. J Cell Biol 1993; 121:855-865. 44. Rogers GE, Taylor LD. The enzymic derivation of citrulline residues from arginine residues in situ during the biosynthesis of hair proteins that are cross-linked by isopeptide bonds. In: Friedman M, ed. Protein Crosslinking. Vol Part A. New York, London: Plenum Press, 1977:283-294. 45. Rogers GE, Winter B, McLaughlan C et al. Peptidylarginine deiminase of the hair follicle: Characterization, localization and function in keratinizing tissues. J Invest Dermatol 1997; 108:700-707. 46. Peterson LL, Zettergren JG, Wuepper KD. Biochemistry of transglutaminases and crosslinking in the skin. J Invest Dermatol 1983; 81:95S-100S. 47. Coulombe PA, Kopan R, Fuchs E. Expression of keratin K14 in the epidermis and hair follicle: Insights into complex programs of differentiation. J Cell Biol 1989; 109:2295-2312. 48. Panteleyev AA, Paus R, Wanner R et al. Keratin 17 gene expression during the murine hair cycle. J Invest Dermatol 1997; 108:324-329. 49. Wilson PA, Short BF. Cell proliferation and cortical cell production in relation to wool growth. Aust J Biol Sci 1979; 32:317-327. 50. Greenwald I, Rubin GM. Making a difference: The role of cell-cell interactions in establishing separate identities for equivalent cells. Cell 1992; 68:271-281. 51. Kopan R, Weintraub H. Mouse notch: Expression in hair follicle correlates with cell fate determination. J Cell Biol 1993; 121:631-641. 52. Kam E, Hodgins MB. Communication compartments in hair follicles and their implication in differentiative control. Development 1992; 114:389-393. 53. Black JL, Reis PJ. Speculation on the control of nutrient partition between wool growth and other body functions. In: Black JL, Reis PJ, eds. Physiological and Environmental Limitations to Wool Growth. Armidale, NSW: University of New England Publishing Unit, 1979:269-293. 54. Downes AM, Chapman RE, Till AR et al. Proliferative cycle and fate of cell nuclei in wool follicles. Nature 1966; 212:477-479. 55. Kalbe J, Kuropka R, Meyer-Stork LS et al. Isolation and characterization of high-molecular mass DNA from hair shafts. Biologische Chemie Hoppe-Seiler 1988; 369:413-416. 56. Schreiber A, Amtmann E, Storch V et al. The extraction of high-molecular mass DNA from hair shafts. FEBS Letters 1988; 230:209-211. 57. Rogers GE. Newer findings on the enzymes and proteins of hair follicles. Ann N Y Acad Sci 1959; 83:408-428. 58. Chapman RE, Ward KA. Histological and biochemical features of the wool fiber and follicle. In: Black JL, Reis J, eds. Physiological and Environmental Limitations to Wool Growth. Armidale, New South Wales, Australia: The University of New England Publishing Unit, 1979:193-208. 59. Mercer JFB, Livingston J, Hall B et al. Isolation of a partial candidate gene for Menkes disease by positional cloning. Nature Genet 1993; 3:20-25.

Late Events and Regulation of Keratinocyte Differentiation in Hair and Feather Follicles

335

60. Takamori K, Thorpe JM, Goldsmith LA. Skin sulfhydryl oxidase. Purification and some properties. Biochim Biophys Acta 1980; 615:309-323. 61. Orwin DFG. An ultrastructural study of the membranes of keratinizing wool follicle cells. J Cell Sci 1972; 11:205-219. 62. Birbeck MSC, Mercer EH. The electron microscopy of the human hair follicle. Part 1. Introduction and the hair cortex. J Biophys Biochem Cytol 1957; 3:203-214. 63. Birbeck MSC, Mercer EH. The electron microscopy of the human hair follicle. Part 2. The hair cuticle. J Biophys Biochem Cytol 1957; 3:215-222. 64. Williams D, Stenn KS. Transection level dictates the pattern of hair follicle sheath growth in vitro. Dev Biol 1994; 165:469-479. 65. Herrling J, Zahn H. Investigations of the cell membrane complex and its modification during industrial processing of wool. Proceedings of the 7th International Wool Textile Research Conference, Tokyo 1985; 1:181-193. 66. Rice RH, Wong VJ, Pinkerton KE. Ultrastructural visualization of cross-linked protein features in epidermal appendages. J Cell Sci 1994; 1985-1992. 67. Allen AK, Ellis J, Rivett DE. The presence of glycoproteins in the cell membrane complex of a variety of keratin fibers. Biochim Biophys Acta 1991; 1074:331-333. 68. Kaytes P, McNab AR, Rea TJ et al. Hair-specific keratins: characterization and expression of a mouse type I keratin gene. J Invest Dermatol 1991; 97:835-842. 69. Wilson BW, Edwards KJ, Sleigh MJ et al. Complete sequence of a type I-microfibrillar keratin gene. Gene 1988; 73:21-31. 70. Compton JG, Ferrera DM, Yu D-W et al. Chromosomal localization of mouse hair keratin genes. Ann N Y Acad Sci 1991; 642:32-43. 71. Rogers MA, Nischt R, Korge B et al. Sequence data and chromosomal localization of human Type I and Type II hair keratin genes. Exp Cell Res 1995; 220:357-362. 72. Hediger R, Ansari HA, Stranzinger GF. Chromosome banding and gene localizations support extensive conservation of chromosome structure between cattle and sheep. Cytogenet Cell Genet 1991; 57:127-134. 73. Yoon S-J, LeBlanc-Straceski J, Ward D et al. Organization of the human keratin type II gene cluster at 12q13. Genomics 1994; 24:502-508. 74. Rosenberg M, RayChaudhury A, Shows TB et al. A group of type I keratin genes on human chromosome 17: Characterization and expression. Mol Cell Biol 1988; 8:722-736. 75. Romano V, Bosco P, Rocchi M et al. Chromosomal assignments of human type I and type II cytokeratin genes to different chromosomes. Cytogenet Cell Genet 1988; 48:148-151. 76. Waseem A, Gough AC, Spurr NK et al. Localization of the gene for human simple epithelial keratin 18 to chromosome 12 using polymerase chain reaction. Genomics 1990; 7:188-194. 77. Powell BC, Cam GR, Fietz MJ et al. Clustered arrangement of keratin intermediate filament genes. Proc Natl Acad Sci USA 1986; 83:5048-5052. 78. Rogers MA, Winter H, Langbein L et al. Genomic characterization of the human type I cuticular hair keratin hHa2 and identification of an adjacent novel type I hair keratin gene hHa5. J Invest Dermatol 1996; 107:633-638. 79. Savtchenko ES, Tomic M, Ivker R et al. Three parallel llinkage groups of human acidic keratin genes. Genomics 1990; 7:394-407. 80. Milisavljevic V, Freedberg IM, Blumenberg M. Close linkage of the two keratin gene clusters in the human genome. Genomics 1996; 34:134-138. 81. Parsons YM, Piper LR, Cooper DW. Linkage relationships between keratin-associated protein (KRTAP) genes and growth hormone in sheep. Genomics 1994; 20:500-502. 82. Nadeau JH, Compton JG, Giguere V et al. Close linkage of retinoic acid receptor genes with homeobox- and keratin-encoding genes on paralogous segments of mouse chromosomes 11 and 15. Mamm Genome 1992; 3:202-208. 83. Hart CP, Compton JG, Langley SH et al. Genetic linkage analysis of the murine developmental mutant velvet coat (Ve) and the distal chromosome 15 developmental genes Hox3.1, Rar-γ, Wnt-1 and Krt-2. J Exptl Zool 1992; 263:83-95.

336

Molecular Basis of Epithelial Appendage Morphogenesis

84. MacKinnon PJ, Powell BC, Rogers GE et al. An ultrahigh-sulfur keratin gene of the human hair cuticle is located at 11q13 and cross-hybridizes with sequences at 11p15. Mamm Genome 1991; 1:53-56. 85. Wood NJ, Phua SH, Crawford AM. A dinucleotide repeat polymorphism at the glycineand tyrosine-rich keratin locus in sheep. Anim Genet 1992; 23:391. 86. French PW, Hewish DR. Localization of low-sulfur keratin proteins in the wool follicle using monoclonal antibodies. J Cell Biol 1986; 102:1412-1418. 87. Heid H, Werner WE, Franke WW. The complement of native α-keratin polypeptides of hair-forming cells: A subset of eight polypeptides that differ from epithelial cytokeratins. Differentiation 1986; 32:101-119. 88. Lynch MH, O’Guin WM, Hardy C et al. Acidic and basic nail/hair (“hard”) keratins: Their colocalization in upper cortical and cuticle cells of the human hair follicle and their relationship to “soft” keratins. J Cell Biol 1986; 103:2593-2606. 89. Kemp DJ, Rogers GE. Immunological and immunofluorescent studies on keratins of the hair follicle. J Cell Sci 1970; 7:273-283. 90. O’Guin WM, Dhouailly D, Manabe M et al. Specific keratins and their associated proteins as markers for hair follicle differentiation. In: Rogers GE, Reis PJ, Ward KA, Marshall RC, eds. The Biology of Wool and Hair. London, New York: Chapman and Hall, 1989:37-50. 91. Kopan R, Fuchs E. A new look into an old problem: Keratins as tools to investigate determination, morphogenesis and differentiation in skin. Genes Dev 1989; 3:1-15. 92. Tobiasch E, Schweizer J, Winter H. Structure and site of expression of a murine type II hair keratin. Mol Biol Rep 1992; 16:39-47. 93. Tobiasch E, Winter H, Schweizer J. Structural features and sites of expression of a new 65kD and 48kD hair-related keratin pair associated with a special type of parakeratotic epithelial differentiation. Differentiation 1992; 50:163-178. 94. Rogers MA, Langbein L, Praetzel S et al. Sequences and differential expression of three novel human type II hair keratins. Differentiation 1997; 61:187-194. 95. Powell BC, Arthur J, Nesci A. Characterization of a gene encoding a cysteine-rich keratin associated protein synthesized late in rabbit hair follicle differentiation. Differentiation 1995; 58:227-232. 96. McNab AR, Andrus P, Wagner TE et al. Hair-specific expression of chloramphenicol acteyltransferase in transgenic mice under the control of an ultra-high-sulfur keratin promoter. Proc Natl Acad Sci USA 1990; 87:6848-6852. 97. Huh N-h, Kashiwagi M, Konishi C et al. Isolation and characterization of a novel hair follicle-specific gene, Hacl-1. J Invest Dermatol 1994; 102:716-720. 98. Fraser RDB, MacRae TP. The fine structure of keratin fibers. In: Breuer MM, ed. Milton Harris: Chemist, Innovator and Entrepreneur. Washington, D.C., USA: American Chemical Society, 1982:119-137. 99. Zhou P, Byrne C, Jacobs J et al. Lymphoid enhancer factor 1 directs hair follicle patterning and epithelial cell fate. Genes Dev 1995; 9:570-583. 100. Powell BC, Nesci A, Rogers GE. Regulation of keratin gene expression in hair follicle differentiation. Ann N Y Acad Sci 1991; 642:1-20. 101. Giese K, Cox J, Grosschedl R. The HMG domain of lymphoid enhancer factor 1 bends DNA and facilitates assembly of functional nucleoprotein structures. Cell 1992; 69:185-195. 102. Giese K, Kingsley C, Kirshner JR et al. Assembly and function of a TCR enhancer complex is dependent on Lef1-induced DNA bending and multiple protein-protein interactions. Genes Dev 1995; 9:995-1008. 103. Sheridan PL, Sheline CT, Cannon K et al. Activation of the HIV-1 enhancer by the Lef1 HMG protein on nucleosome-assembled DNA in vitro. Genes Dev 1995; 9:2090-2104. 104. van Genderen C, Okamura RM, Farinas I et al. Development of several organs that require inductive epithelial-mesenchymal interactions is impaired in Lef1-deficient mice. Genes Dev 1994; 8:2691-2703. 105. Dhouailly D, Xu C, Manabe M et al. Expression of hair-related keratins in a soft epithelium: Subpopulations human and mouse dorsal tongue keratinocytes express keratin markers for hair-, skin- and esophogeal-types of differentiation. Exper Cell Res 1989; 181:141-158.

Late Events and Regulation of Keratinocyte Differentiation in Hair and Feather Follicles

337

106. Kitahara T, Ogawa H. Coexpression of keratins characteristic of skin and hair differentiation in nail cells. J Invest Dermatol 1993; 100:171-175. 107. Stellmach V, Leask A, Fuchs E. Retinoid-mediated transcriptional regulation of keratin genes in human epidermal and squamous cell carcinoma cells. Proc Natl Acad Sci USA 1991; 88:4582-4586. 108. Dlugosz AA, Yuspa SH. Coordinate changes in gene expression which mark the spinous to granular cell transition in epidermis are regulated by protein kinase C. J Cell Biol 1993; 120:217-225. 109. Fuchs E, Byrne C. The epidermis: Rising to the surface. Curr Opin Genet Dev 1994; 4:725-736. 110. Lu B, Rothnagel JA, Longley MA et al. Differentiation-specific expression of human keratin 1 is mediated by a composite AP-1/steroid hormone element. J Biol Chem 1994; 269:7443-7449. 111. Jiang C-K, Tomic-Canic M, Lucas DJ et al. TGF-β promotes the basal phenotype of epidermal keratinocytes: transcriptional induction of K#5 and K#14 keratin genes. Growth Factors 1995; 12:87-97. 112. Byrne C, Tainsky M, Fuchs E. Programming gene expression in developing epidermis. Development 1994; 120:2369-2383. 113. Byrne C, Fuchs E. Probing keratinocyte and differentiation specificity of the human K5 promoter in vitro and in transgenic mice. Mol Cell Biol 1993; 13:3176-3190. 114. Leask A, Byrne C, Fuchs E. Transcription factor AP2 and its role in epidermal-specific gene expression. Proc Natl Acad Sci USA 1991; 88:7948-7952. 115. Leask A, Rosenberg M, Vassar R et al. Regulation of a human epidermal keratin gene: Sequences and nuclear factors involved in keratinocyte-specific transcription. Genes Dev 1990; 4:1985-1998. 116. Magnaldo T, Bernerd F, Freedberg IM et al. Transcriptional regulators of expression of K16, the disease-associated keratin. DNA Cell Biol 1993; 12:911-923. 117. Milisavljevic V, Freedberg IM, Blumenberg M. Characterization of nuclear protein binding sites in the promoter of keratin K17 gene. DNA Cell Biol 1996; 15:65-74. 118. Navarro JM, Casatorres J, Jorcano JL. Elements controlling the expression and induction of the skin hyperproliferation-associated keratin K6. J Biol Chem 1995; 270:21362-21367. 119. Tomic-Canic M, Day D, Samuels HH et al. Novel regulation of keratin gene expression by thyroid hormone and retinoid receptors. J Biol Chem 1996; 271:1416-1423. 120. Casatorres J, Navarro JM, Blessing M et al. Analysis of the control of expression and tissue specificity of the keratin 5 gene, characteristic of basal keratinocytes. J Biol Chem 1994; 269:20489-20496. 121. Huff CA, Yuspa SH, Rosenthal D. Identification of control elements 3' to the human keratin 1 gene that regulate cell type and differentiation-specific expression. J Biol Chem 1993; 268:377-384. 122. de Viragh PA, Huber M, Hohl D. Involucrin mRNA is more abundant in human hair follicles than normal epidermis. J Invest Dermatol 1994; 103:815-819. 123. Takahashi H, Iizuka H. Analysis of the 5'-upstream promoter region of the human involucrin gene: activation by 12-O-tetradecanoylphorbol-13-acetate. J Invest Dermatol 1993; 100:10-15. 124. Welter lF, Crish JF, Agarwal C et al. Fos-related antigen (Fra-1), junB, and junD activate human involucrin promoter transcription by binding to proximal and distal AP1 sites to mediate phorbol ester effects on promoter activity. J Biol Chem 1995; 270:12614-12622. 125. LaPres JJ, Hudson LG. Identification of a functional determinant of differentiation-dependent expression in the involucrin gene. J Biol Chem 1996; 271:23154-23160. 126. Welter JF, Gali H, Crish JF et al. Regulation of human involucrin promoter activity by POU domain proteins. J Biol Chem 1996; 271:14727-14733. 127. Faus I, Hsu H-J, Fuchs E. Oct-6: A regulator of keratinocyte gene expression in stratified squamous epithelia. Mol Cell Biol 1994; 14:3263-3275. 128. Lee J-H, Jang S-I, Yang J-M et al. The proximal promoter of the human transglutaminase 3 gene. J Biol Chem 1996; 271:4561-4568.

338

Molecular Basis of Epithelial Appendage Morphogenesis

129. Maroulakou JG, Papas TS, Green JE. Differential expression of ets-1 and ets-2 protooncogenes during murine embryogenesis. Oncogene 1994; 9:1551-1565. 130. Fisher C, Byers MR, Iadrola MJ et al. Patterns of epithelial expression of Fos protein suggest an important role in the transition from viable to cornified cell during keratinization. Development 1991; 111:253-258. 131. Andersen B, Schonemann MD, Flynn SE et al. Skn-1a and Skn-1i: Two functionally distinct Oct-2-related factors expressed in epidermis. Science 1993; 260:78-82. 132. Sturm RA, Herr W. The POU domain is a bipartite DNA-binding structure. Nature 1988; 336:601-604. 133. Gregg K, Rogers GE. Feather keratin: Composition, structure and biogenesis. In: BereiterHahn J, Matoltsy AG, Richards KS, eds. Biology of the Integument. Vol 2. Berlin: SpringerVerlag, 1986:666-694. 134. Gregg K, Wilton SD, Rogers GE et al. Avian keratin genes: Organization and evolutionary relationships. In: Nagely P, Linnane AW, Pateman JA, eds. Manipulation and Expression of Genes in Eukaryotes. Sydney: Academic Press, 1983:65-72. 135. Presland RB, Gregg K, Molloy PL et al. Avian keratin genes I. A molecular analysis of the structure and expression of a group of feather keratin genes. J Mol Biol 1989; 209:549-559. 136. Whitbread LA, Gregg K, Rogers GE. The structure and expression of a gene encoding chick claw keratin. Gene 1991; 101:223-229. 137. Gregg K, Wilton SD, Parry DAD. A comparison of genomic coding sequences for feather and scale keratins: Structural and evolutionary implications. EMBO J 1984; 3:175-178. 138. Presland RB, Whitbread LA, Rogers GE. Avian keratin genes II. Chromosomal arrangement and close linkage of three gene families. J Mol Biol 1989; 209:561-576. 139. Kemp DJ, Dyer PY, Rogers GE. Keratin synthesis during development of the embryonic chick feather. J Cell Biol 1974; 62:114-132. 140. Rogers GE. Genes for hair and avian keratins. Ann N Y Acad Sci 1985; 455:403-425. 141. Koltunow AM, Gregg K, Rogers GE. Intron sequences modulate feather keratin gene transcription in Xenopus oocytes. Nucl Acids Res 1986; 14:6375-6392. 142. Walker ID, Rogers GE. Differentiation in avian keratinocytes: the properties of the proteins of the chick down feather. Eur J Biochem 1976; 69:329-339. 143. Barnes GL, Sawyer RH. Histidine-rich protein B of embryonic feathers is present in the transient embryonic layers of scutate scales. J Exptl Zool 1995; 271:307-314.

Part V

Models

CHAPTER 17

Epithelial Morphogenesis: A Physico-Evolutionary Interpretation Stuart A. Newman

Introduction

T

he molding of living tissues that occurs during development, regeneration, wound healing, and various pathological processes is referred to as morphogenesis. During morphogenetic events tissue masses may disperse, form internal foci of cell condensation, lengthen or shorten, or acquire lumens. They can also form sheets which may invaginate or evaginate, or develop one or more internal boundaries across which cell mixing is selective or prohibited. Such compartmentalized tissues can physically separate, or remain attached, where they may engulf, or become engulfed by one another. The outcomes of these processes are the various body plans and organ forms characteristic of metazoan organisms, as well as tumors, abnormal polyps and fibrotic lesions. While mechanisms of morphogenesis, like other biological processes, are typically studied by attempting to isolate single determining factors while holding everything else constant, the interactive and regulative nature of developing systems frequently limits the causal information that can be obtained by such strategies. For example, the apical ectoderm of the developing vertebrate limb bud forms a raised ridge over the bud’s distal margin.1 How the ridge forms involves mesodermal effects on the ectoderm, which in turn elicit ectodermal factors which act on the mesoderm.2 Together the two tissues produce an atypical basal lamina at the limb bud margin3-5 which appears to influence the ectodermal cell shape. Exactly how many reciprocal cycles of interaction are required to achieve the end point, and in which tissue the causal chain is initiated, are not clear. It must also be recognized that this morphogenetic event is a product of evolution and, during the course of its attaining its contemporary means of generation, the timing of expression of key genes may been reversed from the order in which their influence was first exerted phylogenetically. This phenomenon, referred to as “heterochrony,”6 can clearly confound experimental assignment of cause and effect. Similarly, whereas the “knocking-out” of a specific gene from an animal’s germ line would seem to be an elegant and straightforward way of assessing its developmental and physiological roles, the readjustment of gene expression and gene product function in the resulting organism can often thwart the interpretation of such experiments. Multicellular organisms first arose as early as 600 million years ago.7 By approximately 540 million years ago, at the end of the “Cambrian explosion,” virtually all the “bauplans” or body types seen in modern organisms already existed.8-10 The original multicellular forms were established with cells that were metabolically and structurally sophisticated—the first single celled organisms appeared two or perhaps three billion years earlier. There is no reason, Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

342

Molecular Basis of Epithelial Appendage Morphogenesis

however, to assume that the earliest metazoa were morphogenetically sophisticated—that they generated complex forms using the baroque, hierarchical molecular machinery that guides such morphogenesis in modern organisms.11 In particular, the defining characteristic of a multicellular organism is simply the existence of a mechanism of adhesion, whereby cells may remain attached to one another after they divide. The precise chemical or physical nature of the adhesive interaction is immaterial, as long as it serves to keep the organism’s cells from entirely dispersing. Therefore the appearance of simple multicellular forms in the fossil record may have been a relatively straightforward matter, and was certainly not dependent on the evolution of any complex developmental schemes. The advent of a cell-cell adhesion mechanism early in the history of multicellular life need not even have required molecular evolution, in the sense of gene sequence change. Some adhesion processes widely used in modern-day organisms, such as those dependent on cadherins, require the availability of calcium ions; in their absence the corresponding cell surface proteins do not mediate cell attachment.12 It is thus conceivable that a simple change in the ionic content of the watery environment of single-celled organisms could have given one or more surface proteins a new function, giving rise to simple metazoan forms by fiat.13 Once one or several adhesive mechanisms were in place, other more complex morphological consequences would have inevitably followed, simply by virtue of “metabolic noise” and physical laws. Cells with different amounts of adhesion molecules on their surfaces, for example, tend not to remain intermixed, but sort out into islands of more cohesive cells within lakes composed of their less cohesive neighbors. Eventually, by random cell movement, the islands will coalesce and multilayered structures will form.14,15 Thus, somewhat counterintuitively, lax regulation of the abundance of adhesion proteins, in conjunction with physical inevitabilities, could have led to novel, but stereotypical, organismal forms. The underlying assumption of the discussion that follows is that much of organismal form originated early in the history of multicellular life by the action of such inevitable or “generic” physical processes on multicellular aggregates (see refs. 16-18 and below). When excitable media (see below) are involved, some of these generic processes can result in what is termed “self-organization,” but the physical effects to which I refer include passive deformations as well, and are therefore of an even broader nature. After the generation of a wide array of forms by interaction of early multicellular organisms with the physical environment, “stabilizing evolution”19 would have led to the “locking-in” of a subset of functionally viable types. In particular, natural selection for organisms that were less subject to the vagaries of the physical environment in their generation of their forms would have resulted in the accumulation of molecular processes with stabilizing and reinforcing roles. Such processes, in turn, would have complemented or superseded purely physical determinants of form. As organisms acquired these “overdetermining” mechanisms, their individual ontogenies would have become more reliably programmed, and they would have thus increasingly bred “true to type.” In effect, phenotypes, which originally would have been only loosely tied to particular genotypes because of the active role of physical externalities in generating form, would over the course of evolution have become more rigidly determined by genotypes.18 This, of course, is the general character of contemporary organisms. As noted, modern organisms have bodies and organs whose morphologies are arrived at by profoundly complex, highly integrated, parallel and redundant means. I suggest that any causal analysis of such complex systems must incorporate an understanding of the physically-based morphogenetic mechanisms that would have prevailed at the earlier stages of the evolution of metazoan form. Although a morphogenetic event in any highly evolved organism can only be fully understood by analyzing all the physical and biochemical inter-

Epithelial Morphogenesis

343

actions that contribute to its realization, over the course of evolution each of these interactions must contribute to different extents to the broad definition of the form in question, or to its fine tuning or stabilization. The view presented here suggests that it is useful to frame such an analysis in terms of the plausible evolutionary history of that morphogenetic event, and the generic physical processes that may have contributed, or continue to contribute, to it.18,20 Only then can the roles of more recently acquired mechanisms of reinforcement and stabilization be understood. Thus, both physics and evolution, two areas usually excluded from the causal analysis of development, are proposed to be absolutely central to this undertaking. In keeping with the theme of this book, the emphasis of this chapter will be on generic morphogenetic mechanisms as they pertain to epithelial tissues, but many of the processes discussed apply equally to mesenchymal and other connective tissues.20

The Earliest Metazoa: Excitable Soft Matter Living tissues, like clay, rubber, lava, and jelly, are viscoelastic or elasticoviscous physical materials, and as such are subject to a set of physical processes that characterize all such substances. The material properties of mature tissues derive partly from the cells that make them up, and partly from the extracellular matrices they produce. Mature epithelioid tissues* contain cells firmly attached to one another, and therefore have rheological properties (including a significant elasticity) akin to those of cytoplasm. The physical properties of mature connective tissues tend to reflect those of their abundant extracellular matrices, which can range from liquid, to gel-like, to solid. In contrast, morphogenetically active epithelioid tissues (i.e., those that participate in embryonic development, regeneration, and neoplasia) contain cells that readily rearrange and slip past one another. The result of this is that these tissues exhibit properties of fluidity, viscosity, and surface tension that are characteristic of liquids.21,22 And because the matrices of morphogenetically active connective tissue are devoid of mineral, and relatively poor in structured fibrous materials, local rearrangement occurs, and these tissues also exhibit liquid-like properties.20 The physicist P.G. De Gennes refers to semi-solid materials such as clays, putties, and polymer melts, as “soft matter”, a category of substance governed by a characteristic set of physical laws.23 The properties of such materials include the capacity to flow, exhibit surface tension, and separate into immiscible phases. A hallmark of soft matter is its capacity to dissipate mechanical stresses through viscous behavior; elastic matter, in contrast, can store and yield back mechanical energy. The evolutionary appearance of multicellular aggregates consisting of freely rearrangeable cells brought organisms for the first time into the realm of soft matter. In addition, these entities were of a spatial scale on which molecular diffusion was no longer instantaneous. Morphogenetically active tissues in modern day organisms indeed frequently behave as if they were composed of soft matter, but it is clear that subsequent evolution has added to the repertoire of tissues the capacity to resist and even oppose passive and externally imposed stresses. While attempts to analyze morphogenetic events in a contemporary organism in terms of the physics of soft matter may entail an unacceptable degree of oversimplification, it is reasonable to assume that more ancient tissues were indeed governed by such laws (see below). Physical scientists have also characterized the behavior of nonliving substances that actively respond to their environment (certain spatially distributed chemical reactors, and networks of coupled electrical elements, are only two such examples), and term this category *Following standard usage, I will refer to cells that adhere by means of direct links to one another as epithelioid. The term epithelial will be reserved for those directly attached cells that have a nonuniform distribution of adhesive molecules on their surfaces, so that they constitute tissues with functionally distinct surfaces, such as skin, kidney tubules, or blood vessels.

344

Molecular Basis of Epithelial Appendage Morphogenesis

of materials “excitable media.”24-27 The fact that even the earliest multicellular organisms would have been composed of chemically active cells, with motile capabilities and the ability to respond to their microenvironments by exhibiting contractile activity and expressing new arrays of gene products, guarantees that from the start metazoa were in this category. Soft matter and excitable media are overlapping but noncoincident categories. In particular, the examples of excitable media given above are not soft matter, nor are all tissues. As discussed previously, most mature tissues, which contain strong linkages between cells, or fibrous or mineralized extracellular matrices, can be considered elastic or inelastic solids, and there is no guarantee that these physical characteristics, which are in the repertoire of all modern tissues, will remain at the sidelines during any given morphogenetic process (see, for example, ref. 28). Morphogenesis in modern-day organisms, then, involves a multiplicity of active and passive physical and chemical processes, reflecting the material complexity of tissues. This complexity can lead to enormous difficulties in attempting to assign pathways of cause and effect during morphogenesis. But because the cells of the earliest metazoa would have lacked the means to establish strong, long-lived connections (tight, anchoring, or gap junctions; rigid matrices) between their surfaces, we can assume that the parcels of tissue that constituted the most primitive metazoa were in the “liquid” state assumed by any condensed material whose subunits (cells, in this case) are independently mobile. These cells, whatever chemical, electrical or contractile excitability they may have been capable of, could have influenced global tissue form only as local responders and actors (and perhaps as sources of diffusible chemical signals), but not, as in many modern situations, as members of a mechanically interlinked lattice. We are thus led to the conclusion that the earliest metazoan organisms consisted of excitable soft matter. This category of material, while hardly simple, has properties that can be analyzed and predicted by standard physical theories and methods. In the following sections I will explore the morphogenetic properties of “fluid epithelia,” as they were presumed to exist prior to the evolution of mechanisms of molecular overdetermination. It is my contention that these properties can provide a taxonomy of the forms assumed by the epithelioid and epithelial tissues of modern organisms. I will also provide a few examples of how these ancient generic properties were recruited to different effect in distinct animal lineages. The specific cellular and molecular mechanisms by which such epithelial forms are attained in their modern embodiments are amply covered in other chapters in this book.

Consequences of Differential Adhesion in Fluid Epithelia Compartment Formation At that point during the history of metazoan organisms when the strength or specificity of intercellular adhesive bonds became subject to modulation, a new class of morphogenetic processes was established: compartment formation. This is a phenomenon by which distinct spatial domains are established within a single tissue, with no interchange or mixing of cells across the common boundary.29,30 It has been demonstrated on both theoretical and experimental grounds that differential adhesion within a tissue mass can lead to the establishment of boundaries across which cells fail to mix.14,15,31,32 What is observed is similar to what happens when two immiscible liquids, such as oil and water, are poured into the same container. As long as the molecules that make up one of the liquids have a greater binding affinity for one another than they do for the molecules of the other liquid, “phase separation” will take place. This takes the form of an interface within the common fluid mass that neither type of molecule will cross, though they may move within their respective liquids by random Brownian motion. Eukaryotic cells are too large to move around by Brown-

Epithelial Morphogenesis

345

Fig. 17.1. Illustration of behavior of intermixed cells and corresponding tissue fragments in the case where the two cell populations are differentially adhesive. The cell mixture will sort out as the more adhesive cells establish more stable bonds with one another than with cells of the other population. Random motion leads to the formation of cohesive islands of these cells, and these will ultimately coalesce into a separate tissue phase, or compartment. Fragments of tissue consisting of the differentially adhesive cell populations will, by spreading, tend to the same equilibrium configuration as the sorted mixture. The boundary between the compartments will have greater curvature with greater disparity of adhesive strengths, with total engulfment of the more cohesive compartment being the extreme case. (Figure adapted from Steinberg,14 1978, with modifications).

ian motion, but they can randomly perambulate through tissues by means of motile forces generated within their cytoplasm. When sufficient differential adhesion exists between two cell populations, not only will each type of cell keep to its own side of the interface, but when dissociated and randomly mixed, the two populations will “sort out”, much like a shaken mixture of oil and vinegar, and for the same thermodynamic reasons (Fig. 17.1). It is important to recognize that whereas compartments in any contemporary tissue system are typically allocated with precision by spatially distributed signals based on molecular gradients (see the section, Pattern Formation in Excitable Epithelia, below), even random assignment of cells to distinct adhesive states can result in a compartmentalized tissue. This is because the sorting-out process will generally bring the cells of similar adhesive state to one side or another of a common boundary (although metastable intermediate states in which the more cohesive population is divided into islands may also persist).33 Thus compartmentalization could have arisen rather early in multicellular evolution, and only later come under the control of biochemically sophisticated regulatory processes. The morphogenetic consequences of compartment formation may be subtle or dramatic. If the adhesive differential is small, the boundary between the two compartments will be relatively straight. If the differential is much greater, one tissue will thereby be more cohesive than the other. It will tend to minimize its surface area, while the less cohesive tissue spreads around, or engulfs, it. Any intermediate degree of differential adhesion will lead to interfaces with various degrees of curvature.14 No matter how straight or curved the interface, the morphogenetic outcome of differential adhesion will be most striking if the cell types that abide by the common boundary are recognizably different from one another. When compartments form in the course of normal development, however, the boundary may be covert to begin with, with the cells in the different compartments becoming structurally or functionally distinguishable only later. Even when the cell types of the cohesively different tissues cannot readily be distinguished, the generation of internal interfacial tensions can produce a multilayered structure where

346

Molecular Basis of Epithelial Appendage Morphogenesis

only a single layer or simple mass existed before, and this can lead to alterations in the shape of a tissue primordium. Whether or not any specific compartmentalization event in a modern-day organism— the formation of anterior and posterior compartments in the Drosophila wing,29,30 for example—is due purely to differential adhesion, or whether other, or additional, barriers to cell mixing are introduced during the course of development, can only be determined experimentally. What the preceding discussion has shown is that differential adhesion is sufficient to cause compartmentalization of an epithelioid tissue, and that it is probably the simplest way that this phenomenon could arise in a multicellular tissue governed by the laws of soft matter.

Cell Polarity and Lumen Formation Epithelioid cells can in principle have uniform adhesive properties around their entire surfaces; the blastomeres of the early mammalian embryo are a case in point. Many epithelioid, and all epithelial cell types, however, are polarized in the expression of several functions, notably adhesion.34 The targeting of adhesive molecules, or anti-adhesive molecules, to specific regions of the cell surface can have dramatic consequences. A tissue mass consisting of motile epithelioid cells that are non-adhesive over portions of their surfaces would readily develop cavities or lumens (Fig. 17.2). If such spaces were to come to adjoin one another, as a result of random cell movement, they would readily fuse. Lumen formation may therefore have originated as a simple consequence of differential adhesion in cells that express adhesive properties in a polarized fashion. In a recent study, for example, the met oncogene, which encodes a cytokine receptor, was ectopically expressed in epithelioid mammary carcinoma cells, which do not normally form lumens.35 When the cells were then stimulated with the ligand HGF/SF, the tumor tissue formed lumens, and Met protein was specifically localized over the apical region of each cell bordering a lumen. A simple interpretation (although one that has not yet been tested directly) is that the polarized localization of Met renders a portion of each expressing cell nonadhesive, and by cell rearrangement this becomes the region bordering the lumen. The notion that lumen formation is a consequence of a delicate balance of adhesive interactions between cells of the same and different types, and/or their extracellular substrata, is supported by experimental studies and genetic analyses of various developmental and pathological conditions. Mammary epithelial cells when grown on tissue culture plastic in the absence of ECM adopt a flat “cobblestone” appearance. In the presence of laminin, however, they round up and cluster and, depending on the culture conditions, may form hollow, alveolar structures with well-defined apical and basal surfaces.36 Salivary gland and lung epithelial cells similarly undergo, or fail to undergo, branching tubular or alveolar morphogenesis depending on specific cell-ECM interactions (reviewed in ref. 37). Recent studies of human autosomal dominant polycystic kidney disease (ADPKD) indicate that the formation of cysts, rather than tubules, in the kidneys of severely affected patients, involves the expression of mutated forms of polycystin,38 a putative integral membrane glycoprotein that is thought to mediate cell-cell or cell-matrix interactions of polarized kidney epithelial cells.39 Depending on the taxon, cell polarity could have arisen before or after the evolutionary event that led to multicellularity. It is important to recognize, however, that lumen or cavity formation would have been an inevitable physical consequence of the conjunction of these two properties. The evolution of the epithelioid-epithelial transition would have laid the basis for various cavitation processes in the tissues of modern organisms, although this would have required that the transition come under more stringent ontogenetic control.

Epithelial Morphogenesis

347

Fig. 17.2. Schematic view of formation of a lumen or internal cavity by differential adhesion in an epithelioid tissue consisting of polarized cells. In the original state (left) the cells are uniformly adhesive, and make contacts around their entire peripheries. Upon expression of an anti-adhesive protein in a polarized fashion in a random subpopulation of cells (center), and random movement of the cells throughout the mass, bonds between adhesive surfaces are energetically favored over those between adhesive and nonadhesive surfaces, resulting in lumen formation (right). (Figure adapted from Newman and Tomasek,20 1996, with modifications).

A further consequence of the polar expression of adhesive function would have been the establishment of cell sheets with two free surfaces. This would have required a more sophisticated regulation of the targeting of homotypic adhesive molecules, which confined their expression to lateral cell boundaries. Of course, apposition of a simply polarized tissue (i.e., one in which adhesive molecules were excluded from only one portion of the cell) with a second tissue which adhered to the cells of the first more strongly than they adhered to one another, would inevitably lead to the formation of a cell sheet with one free and one attached surface, as is seen in skin and other epithelial-mesenchymal units. Other cellular mechanisms, such as the contraction of apical actin filaments in a group of cells in a localized domain of an epithelial sheet, undoubtedly contribute to, and may even precipitate, lumen formation in contemporary organisms. But this mechanism also requires polarized cells, and in addition requires a global pattern formation system to specify the position of the contracting domain (see section, Shape Change in Epithelial Sheets, below). In the absence of such a system (e.g., in the earliest metazoa, and perhaps in some modern developmental processes like salivary gland or testis morphogenesis, where precise spatial regulation is not required), targeting of adhesive or anti-adhesive molecules to restricted plasma membrane domains, in conjunction with sorting-out, provides the most plausible route to the formation of intratissue cavities or lumens.

Pattern Formation in Excitable Epithelia As we have seen, spatial variations in adhesion within an aggregate readily leads to compartment formation and tissue multilayering, while regional variations in adhesion at the level of the individual cell readily leads to lumen formation. It was suggested that these “spontaneous” behaviors of fluid epithelia were made use of early in the evolution of metazoa, and may also underlie some aspects of epithelial morphogenesis in modern animals. However, if such spatial and regional variations in adhesion were simply a random occurrence, the capacity for organisms to undergo precisely patterned developmental programs in each successive generation would be severely limited. It is significant, therefore, that the identity of multicellular aggregates as macroscopic objects (i.e., of a scale on which diffusion is not instantaneous), and as excitable media (features that up to now we have not made use of in characterizing epithelial behavior) bring a novel set of regionalizing or pattern forming processes into play. These, in turn, may influence local expression of adhesive molecules,

348

Molecular Basis of Epithelial Appendage Morphogenesis

contractility, pigmentation, or other cell properties, giving rise to epithelia, or other tissues, with well-regulated shapes and patterns. The reacting and diffusing chemicals within cells and tissues provide the conditions for several physicochemical processes that can give rise to molecular nonuniformities in space and time. Gradients of chemical composition can be formed by diffusion from a localized source which may be established when an initially isotropic founder cell interacts with a chemically or physically nonuniform environment. The diffusing molecule could be passed directly from cell to cell by transport across membranes,40 or indirectly, through the extracellular space.41 If a nonuniformly distributed chemical came to influence the expression of adhesivity in the cluster of cells produced by this founder, reproducible compartmentalization would result, and as we have seen, this could drive morphogenetic rearrangement in a predictable fashion. Cellular metabolism would have included many coupled chemical reactions even in the pre-metazoan stages of evolution. Such reactions, when present in multicellular aggregates, are capable of becoming organized in time and space. For instance, positive feedback of the production of a chemical species, in the context of an otherwise self-limiting set of reactions, will often give rise to temporal oscillations in the concentration of one or more of the reactants.42-44 Such chemical oscillations, which are well-known experimentally,45-47 can occur in countless different reaction systems, which need only fulfill a set of formal kinetic conditions in order for temporally periodic changes in concentration to occur. If linked to the control of any cellular behavior, such as the expression of pigment, or of adhesion molecules, chemical oscillations in time can provide the basis for tissue properties that are periodic in space (see below). The joint effects of positive autoregulation and cross-inhibition in a chemical reaction system with diffusion of one or more of the chemicals, can lead to an unusual form of spatial organization: stable, nonuniform patterns of concentration of one or more of the molecules (Fig. 17.3). This phenomenon was investigated by Turing,48 who hypothesized it to be “the chemical basis of morphogenesis.” More recently, “Turing patterns” have been demonstrated experimentally in several nonliving chemical systems reacting within semi-solid media,49-51 where they take the form of gradients, stripes, spots, or even spirals52 of chemical concentration. A Turing-type “reaction-diffusion” mechanism has been proposed to generate a prepattern of secreted gene products underlying eyespot patterns on butterfly wings.53 Some of the gene products that organize into this prepattern have been identified, and they are identical to those which cue the Drosophila wing imaginal disk to evaginate into the wing blade.54 By extension, then, an adhesion-based morphogenetic process in the Drosophila wing epithelium may be controlled by the same reaction-diffusion process that in the butterfly controls epithelial cell pigment pattern formation.55 This confirms an important expectation of the view presented here: that pattern forming processes generic to epithelia considered as excitable media can have become linked to the expression of distinct cellular properties (e.g., pigmentation, differential adhesion, contraction (see below)) on independent occasions over the course of evolution.

Segmentation The formation of stripes or spots of pigment in an epithelium, as in the skin of certain mammals or fishes, is a striking consequence of epithelial pattern formation, but has little overtly to do with morphogenesis. If, however, the same types of pattern forming processes became linked to the production of cell adhesion molecules, dramatic morphological consequences could follow. For example, a hexagonal arrangement of peaks of an inductive molecule (readily produced by a Turing-type reaction-diffusion process) could alter adhesive properties in an epithelial sheet, giving rise to the epithelial placodes that provide the primordia for developing feathers.56-58

Epithelial Morphogenesis

349 Fig. 17.3. Graphical representation of chemical wave formation by a Turing-type reaction-diffusion mechanism. It is assumed that two substances, A (solid line) and B (broken line), which influence one another’s synthesis, are produced throughout a row of cells, and that the balance between the rates of synthesis and utilization of A and B causes each substance to attain a spatially uniform steady state (i.e., temporally stationary) concentration profile along the row of cells. Under certain conditions a spatially nonuniform stationary state (e) can be achieved by the growth and stabilization of a random fluctuation (b-d).48 The following conditions are sufficient to bring about this phenomenon: Substance A has a positive effect on the synthesis of both itself and substance B; substance B has an inhibitory effect on the synthesis of A; the diffusion rate of B is greater than that of A. Arrow in (c) indicates the point at which a reduction of the concentration of A to below its uniform steady state level will be initiated, based on the assumptions above. The number of peaks and valleys of A and B that will be in place when the system finally reaches the new spatially nonuniform steady state will depend on reaction and diffusion rates, the size and shape of the spatial domain in which these events are occurring, and the modes of utilization of A and B at the boundaries of the domain. (Figure adapted from Maynard Smith16 1968, with modifications).

350

Molecular Basis of Epithelial Appendage Morphogenesis

Another example, with profound consequences for the organization of body plans, is segmental organization: the repetitive alternation of tissue compartments, understood in the sense of immiscibility across a common boundary (see section, Consequences of Differential Adhesion in Fluid Epithelia, above). Such spatial periodicities can be arrived at developmentally over a period of time, or simultaneously, over a particular tissue domain. For example, in short germ band insects and crustaceans, segmental primordia are added in sequence from a caudal zone of cell proliferation.59,60 In contrast, in long germ band insects, such as Drosophila, a series of “chemical stripes,” consisting of alternating evenly-spaced bands of transcription factors specified by certain “pair-rule” genes in the syncytial embryo, ensure that when cellularization finally takes place the cells of the blastoderm have periodically distributed identities. These covert cell identities are later converted into states of differential adhesivity,61 and overt morphological segments form as a consequence. While it seems puzzling that evolutionarily related insects such as beetles (short germ band) and fruit flies (long germ band) have such different modes of segment formation, consideration of tissues as excitable media can unify the understanding of these phenomena.17 First it is necessary to consider how a temporally varying chemical oscillator can organize a developing tissue into alternating immiscible blocks. Let us assume, as an example, that there is a localized embryonic zone containing a synchronized population of cells which divide at regular intervals. We will also assume that the number of adhesive molecules on the surfaces of these cells is set at the time of mitosis, as a function of the cellular concentration of a regulatory molecule, R, and that each cell retains its “adhesive state” during its lifetime. Now if the concentration of R were to oscillate with time, and if the period of this oscillation were the same as that of the cell cycle, each cell would then have the same adhesive state, and the tissue so generated would have no segmental character. If, in contrast, the period of the R oscillation were different from that of the cell cycle, successive populations of cells would be born with different adhesive states, and the changing phase relation between the cell cycle and adhesivity-regulating oscillators would ensure that adhesive states would recur periodically. An example of how this “temporal phase shift” mechanism can generate segments is shown in Figure 17.4.17 The preceding analysis now permits us to formulate a plausible scenario for how the “temporal” and “spatial” modes of segmentation might have arisen in taxonomically related groups. This depends on the recognition that the same types of chemical systems that permit oscillations to arise can readily form Turing structures if diffusion is added to the dynamic mix.62 And indeed, in the syncytial embryo of long germ band insects such as Drosophila, several of the factors that become organized into the early chemical stripes freely diffuse among the cell nuclei that synthesize their mRNAs. Some of these also positively regulate their own synthesis,63,64 a sine qua non of both chemical oscillators and Turing pattern forming systems (Fig. 17.5). As discussed in earlier sections, there is every reason to expect that once a successful morphological motif is established in a particular taxonomic group, the developmental mechanisms by which it was ontogenetically achieved would undergo stabilizing evolution, becoming more complex at the molecular level. Thus, the observation that striped expression of the Drosophila pair-rule genes often involves multiple promoter elements responsive to preexisting, nonuniformly distributed molecular cues (e.g., maternal and gap gene products)65,66 is not inconsistent with this pattern having originated as a Turing process. While this suggestion must remain hypothetical in the absence of any direct evidence about ancient mechanisms of segmental pattern formation, it should be noted that by the reaction-diffusion mechanism chemical stripe formation is achieved simply, with a minimum of molecular ingredients and physical processes. It therefore represents a more plausible early basis for the striped distribution of pair rule proteins than the alternative that the

Epithelial Morphogenesis

351 Fig. 17.4. Model for the generation of segments in a zone of synchronized cell multiplication, by the temporal oscillation of the concentration of a molecule that regulates cell adhesion. The clock faces represent the phase of the cell cycle C and that of the periodically varying regulatory molecule R. It is assumed in this example that the duration of the cell cycle is three hours, the period of the chemical oscillation is two hours, and that both cycles start together. During the first cell cycle, newly formed cells have an adhesive state specified by the initial value of R (stippling). During the second cell cycle, R is in mid-cycle, and the newly formed cells have a different adhesive state (hatched lines). During the third cell cycle R is again at its initial concentration, and the new cells have the first adhesive state. The assumption of cell synchrony is for simplification of the model; the mechanism would also give rise to segments in a zone of asynchronous cell multiplication with local cell sorting-out. (Figure adapted from Newman,17 1993, with modificaitons).

Fig. 17.5. Schematic representation of possible relationship between two modes of tissue segmentation. Both modes have in common a biochemical circuit that generates a chemical oscillation. One of the oscillating species directly or indirectly regulates the strength or specificity of cell adhesivity. In the mechanism shown on the left, the periodic change in cell adhesivity occurs in a growth zone in which the cell cycle has a different period from the regulatory oscillator; as a result, bands of tissue are sequentially generated with alternating cohesive properties (see Fig. 17.4). In the mechanism shown on the right, one or more of the biochemical species can diffuse, leading to a set of standing waves of concentration of the regulatory molecule by a reaction-diffusion mechanism (see Fig. 17.3). This leads to the simultaneous formation of bands of tissue with alternating cohesive properties. See ref.17 for additional details.

352

Molecular Basis of Epithelial Appendage Morphogenesis

stripes originated independently, each depending on separate and unique promoter-transcription factor interactions. Apart from the questions of how segmental organization originated evolutionarily, and how it is achieved during embryogenesis in any modern organism, it is reasonable to ask whether segmental boundaries in segmented tissues indeed result from alternating adhesive affinities, as hypothesized here. For one segmented epithelioid tissue, the vertebrate hindbrain, which consists of blocks of neuroepithelial cells referred to as rhombomeres, this can be answered in the affirmative. In grafting experiments on the chick embryo hindbrain, it was found that juxtaposition of tissue from any position within adjacent rhombomeres led to the regeneration of a barrier to cell mixing, but juxtaposition of tissue from alternate units did not.67 More detailed analysis of this system revealed a hierarchy of differential adhesive interactions emerging over the course of development that first block out broad regions of nonmixing cells and then refine the rhombomeric pattern.68 Eventually, specializations develop at the rhombomere boundaries that provide mechanical, rather than adhesive, barriers to cell mixing.69 According to the framework presented here, this developmental sequence can productively be interpreted as the evolved refinement of a segmentation process primitively based on the regulation of cell adhesion by chemical oscillations in an excitable neuroepithelium.

Shape Change in Epithelial Sheets While simple compartment formation can occur in a tissue by self-organization resulting from differential adhesion (see section, Consequences of Differential Adhesion in Fluid Epithelia, above), the example of segmentation demonstrates that once a chemical pattern forming system is in place, more complex morphological outcomes are possible. This is because numerous cell activities can now be modulated in a reproducible position-dependent fashion within the cell mass or sheet. Assuming fluid rearrangement of the cells, cell adhesive preference is certainly one of these properties, and this has been considered extensively, above. However, Gierer70 also points to a variety of other cellular activities which can contribute to cell shape change, such as production of intracellular tensile forces by cell surface-cytoskeletal interactions. Like cell adhesion, which is subject to a “minimum energy” principle (reflected in sorting-out behavior, for example), the other processes considered by Gierer (insofar as they are reversible) are subject to analogous quasi-equilibrium principles: Under a given set of environmental constraints the corresponding energies of interaction will also tend to a minimum. Since such properties (referred to as “potentials” or “potential functions”), have this characteristic in common, they can be analyzed collectively, using a general formalism.70 Of course, irreversible changes in tissue organization, which may accompany cell differentiation, for instance, could also contribute to tissue morphogenesis. Such changes would not be analyzable by the pattern formation-cell potential function formalism. However, Gierer makes the reasonable assumption that reversible interactions characterize the ontogenetically earliest (and I would add, with even more likelihood, the phylogenetically earliest) morphogenetic processes, and that many regulatory features seen in early development derive from this property. It is worth reiterating that cell adhesion is unique among such cell potentials in having morphogenetic consequences even in the absence of pattern formation, since cells can sort out into discrete compartments even when adhesive states are assigned randomly. Random assignment of cell shape change, in contrast, would by itself have little or no global effect on the morphology of a cell mass or sheet. When changes in parameters affecting cell potentials are assigned to particular cell populations by a globally-acting pattern-forming mechanism, however, a coordinated alteration in tissue morphology is possible, even if cell rear-

Epithelial Morphogenesis

353

Fig. 17.6. Schematic representation of epithelial morphogenesis by patterned alteration of cell parameters. In (A), a pattern formation process (e.g., a reactiondiffusion system) is activated in a flat epithelial sheet, and marks a subset of cells to undergo alteration of one or more cell “potential functions” (e.g., adhesive strength, cytoskeletal tension, or any other parameter subject to a “minimal energy” principle). In (B), the case is shown in which the alteration changes the cells in the lateral aspect only. Increase in extent of lateral adhesive interaction, or in the cell’s contractile state in the horizontal dimension, leads to the formation of a placode. In (C), the change in cell potential gives rise to a bending moment that destabilizes the flat configuration, leading to evagination or invagination.

rangement is limited. Consider Figure 17.6, in which a patterning process is assumed to designate a subset of cells in a flat sheet for alteration of cell potential (Fig. 17.6A). The resulting changes (which may be in adhesive properties, internal cytoarchitecture, or any other reversible influence on cell shape) might be exerted in the lateral direction alone, whereby a placode would form (Fig. 17.6B). With a different set of external constraints, however, the flatness of the sheet could be destabilized by the generation of a bending moment, and the tissue would evaginate or invaginate (Fig. 17.6C).70 The reversibility that qualifies cell activities to be treated as potential functions need not persist after the initial morphogenetic events have occurred. As noted above, and as discussed by Gierer,70 the mechanical properties of a mature or established tissue, while built on templates generated by the processes described, also depend on long-lived intercellular bonds and matrices and on intracellular reorganization consequent to cell differentiation. An additional morphogenetic mechanism pertaining to epithelia was arrived at by Mittenthal and Mazo in their consideration of the role of elasticity in cell sheets.71 As discussed above, the fundamental assumption in our physical analysis of epithelial morphogenesis is that the independent mobility of cells ensures that the tissues behave as soft matter, with elasticity making only a marginal contribution. Significantly, Mittenthal and Mazo base their analysis on the same general assumption, considering their model epithelial sheets to be entirely liquid in the tissue plane. Based on certain experimental findings, however (e.g., studies on eversion of Drosophila imaginal discs72), they make the further assumption that unlike other two-dimensional liquids such as soap films, epithelia have a globally-distributed elastic component that resists bending outside of the plane of the tissue. Thus they treat the epithelium as a “fluid elastic shell”, fluid-like in its capacity for in-plane rearrangement of cells, but resembling an elastic sheet in bending.71 The basis of the out-of-plane elastic component is suggested by these authors to be the epithelial cell membranes or the extracellular matrix. While recent studies of the determinants of cell shape suggest that the elastic properties of the cell membrane are not significant,73 all known epithelia have basal laminae consisting largely of dense sheets of collagenous protein,74 and these structures are likely to provide resistance to bending of the tissue sheet.

354

Molecular Basis of Epithelial Appendage Morphogenesis

The phenomenon for which Mittenthal and Mazo seek an account is the relationship of width to length—the curvature—of an epithelial tube, such as an arthropod leg. The tube they consider is a series of bands of differentially adhesive cells, generated in their analysis by cell sorting of an initially dispersed mixture. The tissue forms a “target” pattern as a result of the hierarchy of engulfment relations. Then (subject to the stiffening constraint that keeps the sheet planar and elastic) it evaginates into a hollow tube, so as to reduce the contact, and hence the adhesive disparity, between adjacent bands. Thus far, their analysis is equivalent to that of Gierer,70 with adhesivity serving as the potential function, and cell sorting as the pattern-forming mechanism. The authors then derive a mathematical relationship that predicts the tube shape, which minimizes a potential function consisting of both the elastic and adhesive energies. The predictions of this “scaling relationship” are compared with measurements of the different leg segments in Drosophila, and the fit is relatively good. For a discussion of improvements to the model, and the validity of its underlying assumptions (which, like the assumptions in the previous sections of this review, are even more pertinent for phylogenetically ancient than for contemporary forms), the reader is referred to the original paper.71

Conclusion In this review I have sketched out some major morphogenetic processes that pertain to epithelioid and epithelial tissues. These processes are abstracted from actual, contemporary multicellular systems in several important respects. First of all, I have confined my attention to tissue properties that are expected to have prevailed at early stages of multicellular evolution. For example, the assumption of independent cell mobility, which is valid for some,22 but certainly not all, contemporary embryonic tissues, provides the basis for the liquid tissue- or soft matter-based interpretation of their behaviors considered here. In addition, the treatment of tissues as excitable media (clearly a category that even highly evolved modern tissues belong to) emphasizes dynamic processes such as chemical oscillations and reaction-diffusion mechanisms, which, while occasionally present in modern organisms, must also have been displaced to varying extents by more “hard-wired” regulatory processes in the course of evolution. These abstractions have permitted a focus on physical processes which presumably established morphological templates for the subsequent evolution of complexity at the molecular level. Moreover, these assumptions have afforded the possibility of examining relatively simple chains of causation in morphogenesis, which is all but precluded in the analysis of highly evolved systems. The responsiveness of tissues to, and recovery from, perturbation (referred to in physiological systems as “homeostasis”—stability of state, or in developing systems as “homeorheosis”—stability of pathway75), which are among the most striking characteristics distinguishing the living from the nonliving, are de-emphasized in the framework presented here. The justification for this is that these rectifying mechanisms were undoubtedly acquired over the course of long periods of evolution, so that the more malleable and polymorphic systems considered here represent actual tissues of an earlier era. If this hypothesis is valid, then the forms assumed by modern tissues can only be understood by this sort of abstraction.18,76 Secondly, the discussion presented here has largely ignored inductive interactions between distinct tissue types. The most primitive such interactions may have occurred between the distinct epithelioid tissue types that first arose as a consequence of compartmentalization. Once they became compartmentalized, tissues could acquire distinct biochemical identities that extend beyond the adhesive differences that first defined them as separate. They could then begin to influence one another’s fates across their common boundaries, adding further complexity to the pattern-forming processes discussed here. The relation-

Epithelial Morphogenesis

355

ship between compartmentalization and induction is considered by Meinhardt77 who hypothesizes that formation of internal tissue boundaries can provide new interfaces for inductive interactions in the course of development. But this relationship can also be considered evolutionarily, with compartmentalization plausibly providing the original platform for tissue diversification and interaction.18 Any comprehensive understanding of the mechanisms of epithelial morphogenesis must take these later appearing inductive phenomena into account, along with the more primitive generic physical determinants discussed here. Finally, the morphogenetic processes unique to, and indeed generic to, mesenchymal and other connective tissues—local cell condensation, tissue contraction (reviewed in ref. 20), and their inductive and mechanical influences on epithelia over the course of evolution, and in contemporary systems, have been largely left out of the present discussion. While many animal taxa clearly evolved without the participation of a connective tissue lineage, it is implausible that the evolution of epithelial morphogenesis was independent of that of its mesenchymal partner in organisms containing both types of tissue. In particular, we have previously suggested that the extracellular matrix of mesenchymal tissues, which supports the diffusion of peptide growth factors, is a likely biological medium for the realization of Turing-type chemical waves.17,41,78 Thus some periodicities seen in epithelia, such as the feather germs of avian skin,56,79 may depend on the intrinsic pattern forming properties, or “excitability,” of the underlying mesenchymal component. The foregoing discussion has shown that by introducing concepts from the physics of condensed materials and the evolution of developmental mechanisms into the analysis of morphogenesis, it is possible to overcome certain intrinsic circularities in standard attempts to account for tissue pattern and form. But it is also clear that, while the notions presented in this review may be a necessary starting point for a comprehensive causal analysis of the morphogenetic mechanics of contemporary tissue systems, they are only a starting point.

References 1. Saunders JW Jr. The proximo-distal sequence of origin of the parts of the chick wing and the role of the ectoderm. J Exp Zool 1948; 108:363-402. 2. Saunders JW Jr. The experimental analysis of chick limb bud development. In: Ede DA, Hinchliffe JR, Balls M, editors. Vertebrate Limb and Somite Morphogenesis. Cambridge: Cambridge University Press, 1977:1-24. 3. Tomasek JJ, Mazurkiewicz JE, Newman SA. Nonuniform distribution of fibronectin during avian limb development. Dev Biol 1982; 90:118-26. 4. Newman SA, Frisch HL, Perle MA et al. Limb development: Aspects of differentiation, pattern formation and morphogenesis. In: Connolly TG, Brinkley LL, Carlson BM, editors. Morphogenesis and Pattern Formation. New York: Raven Press, 1981:163-78. 5. Critchlow MA, Hinchliffe JR. Immunolocalization of basement membrane components and beta 1 integrin in the chick wing bud identifies specialized properties of the apical ectodermal ridge. Dev Biol 1994; 163:253-69. 6. Gould SJ. Ontogeny and Phylogeny. Cambridge, MA: Harvard University Press, 1977. 7. Conway Morris S. The fossil record and the early evolution of the Metazoa. Nature 1993; 361:219-25. 8. Whittington HB. The Burgess Shale. New Haven: Yale University Press, 1985. 9. Conway Morris S. Burgess shale faunas and the cambrian explosion. Science 1989; 246:339-46. 10. Briggs DEG, Fortey RA, Wills MA. Morphological disparity in the cambrian. Science 1992; 256:1670-3. 11. Nusslein-Volhard C. Gradients that organize embryo development. Sci Am 1996; 275:54-5; 58-61. 12. Takeichi M. Cadherin cell adhesion receptors as a morphogenetic regulator. Science 1991; 251:1451-5.

356

Molecular Basis of Epithelial Appendage Morphogenesis

13. Kazmierczak J, Degens ET. Calcium and the early eukaryotes. Mitt Geol-Palaeont Inst Univ Hamburg 1986; 61:S. 1-20. 14. Steinberg MS. Specific cell ligands and the differential adhesion hypothesis: How do they fit together? In: Garrod DR, editor. Specificity of Embryological Interactions. London: Chapman and Hall, 1978:97-130. 15. Steinberg MS, Takeichi M. Experimental specification of cell sorting, tissue spreading, and specific spatial patterning by quantitative differences in cadherin expression. Proc Natl Acad Sci U S A 1994; 91:206-9. 16. Newman SA, Comper WD. ‘Generic’ physical mechanisms of morphogenesis and pattern formation. Development 1990; 110:1-18. 17. Newman SA. Is segmentation generic? BioEssays 1993; 15:277-83. 18. Newman SA. Generic physical mechanisms of tissue morphogenesis: A common basis for development and evolution. J Evol Biol 1994; 7:467-88. 19. Schmalhausen II. Factors of Evolution. Philadelphia: Blakiston, 1949. Trans, Dordick I. 20. Newman SA, Tomasek JJ. Morphogenesis of connective tissues. In: Comper WD, editor. Extracellular Matrices. v. 2: Molecular Components and Interactions. Reading, U.K.: Harwood Academic Publishers, 1996:335-69. 21. Steinberg MS, Poole TJ. Liquid behavior of embryonic tissues. In: Bellairs R, Curtis ASG, editors. Cell Behavior. Cambridge: Cambridge University Press, 1982:583-607. 22. Foty RA, Pfleger CM, Forgacs G et al. Surface tensions of embryonic tissues predict their mutual envelopment behavior. Development 1996; 122:1611-20. 23. de Gennes PG. Soft matter. Science 1992; 256:495-7. 24. Mikhailov AS. Foundations of Synergetics I. Berlin: Springer-Verlag, 1990. 25. Gerhardt M, Schuster H, Tyson JJ. A cellular automation model of excitable media including curvature and dispersion. Science 1990; 247:1563-6. 26. Starmer CF, Biktashev VN, Romashko DN et al. Vulnerability in an excitable medium: analytical and numerical studies of initiating unidirectional propagation. Biophys J 1993; 65:1775-87. 27. Winfree AT. Persistent tangled vortex rings in generic excitable media. Nature 1994; 371:233-6. 28. Davidson LA, Koehl MA, Keller R et al. How do sea urchins invaginate? Using biomechanics to distinguish between mechanisms of primary invagination. Development 1995; 121:2005-18. 29. Crick FHC, Lawrence PA. Compartments and polyclones in insect development. Science 1975; 189:340-7. 30. Garcia-Bellido A, Ripoll P, Morata G. Developmental compartmentalization in the dorsal mesothoracic disc of Drosophila. Develop Biol 1976; 48:132-47. 31. Armstrong PB. Cell sorting out: The self-assembly of tissues in vitro. Crit Rev Biochem and Mol Biol 1989; 24:119-49. 32. Graner F. Can surface adhesion drive cell-rearrangement? Part I: Biological cell-sorting. J Theor Biol 1993; 164:455-76. 33. Antonelli PL, Rogers TD, Willard MA. Geometry and the exchange principle in cell aggregation kinetics. J Theor Biol 1973; 41:1-21. 34. Rodriguez-Boulan E, Nelson WJ, editors. Epithelial and Neuronal Cell Polarity. Cambridge: Company of Biologists, 1993. 35. Tsarfaty I, Resau JH, Rulong S et al. The met proto-oncogene receptor and lumen formation. Science 1992; 257:1258-61. 36. Li ML, Aggeler J, Farson DA et al. Influence of a reconstituted basement membrane and its components on casein gene expression and secretion in mouse mammary epithelial cells. Proc Natl Acad Sci U S A 1987; 84:136-40. 37. Ashkenas J, Muschler J, Bissell MJ. The extracellular matrix in epithelial biology: Shared molecules and common themes in distant phyla. Develop Biol 1996; 180:433-44. 38. Qian F, Watnick TJ, Onuchic LF et al. The molecular basis of focal cyst formation in human autosomal dominant polycystic kidney disease type I. Cell 1996; 87:979-87.

Epithelial Morphogenesis

357

39. Hughes J, Ward CJ, Peral B et al. The polycystic kidney disease 1 (PKD1) gene encodes a novel protein with multiple cell recognition domains. Nat Genet 1995; 10:151-60. 40. Crick FHC. Diffusion in embryogenesis. Nature 1970; 225:420-2. 41. Newman SA, Frisch HL. Dynamics of skeletal pattern formation in developing chick limb. Science 1979; 205:662-8. 42. Goldbeter A, Lefever R. Dissipative structures for an allosteric model: Application to glycolytic oscillations. Biophys J 1972; 12:1302-15. 43. Norel R, Agur Z. A model for the adjustment of the mitotic clock by cyclin and MPF levels. Science 1991; 251:1076-8. 44. Tyson JJ. Modeling the cell division cycle: cdc2 and cyclin interactions. Proc Nat Acad Sci USA 1991; 88:7328-32. 45. Chance B, Estabrook RW, Ghosh A. Damped sinuosoidal oscillations of cytoplasmic reduced pyridine nucleotide in yeast cells. Proc Nat Acad Sci USA 1964; 51:1244-51. 46. Nurse P. Universal control mechanisms regulating onset of M-phase. Nature 1990; 344:503-8. 47. Hall JC, Rosbash M. Oscillating molecules and how they move circadian clocks across evolutionary boundaries. Proc Natl Acad Sci U S A 1993; 90:5382-3. 48. Turing A. The chemical basis of morphogenesis. Phil Trans Roy Soc Lond B 1952; 237:37-72. 49. Castets V, Dulos E, Boissonade J, DeKepper P. Experimental evidence of a sustained standing Turing-type nonequilibrium chemical pattern. Phys Rev Lett 1990; 64:2953-6. 50. Ouyang Q, Swinney H. Transition from a uniform state to hexagonal and striped Turing patterns. Nature 1991; 352:610-2. 51. Lengyel I, Kadar S, Epstein IR. Transient Turing structures in a gradient-free closed system. Science 1993; 259:493-5. 52. Epstein IR. Spiral waves in chemistry and biology. Science 1991; 252:67. 53. Nijhout HF. The Development and Evolution of Butterfly Wing Patterns. Washington: Smithsonian Inst Press, 1991. 54. Carroll SB, Gates J, Keys DN et al. Pattern formation and eyespot determination in butterfly wings. Science 1994; 265:109-14. 55. Nijhout HF. Genes on the wing. Science 1994; 265:44-5. 56. Sengel P. Morphogenesis of Skin. Cambridge: Cambridge Univ. Press, 1976. 57. Chuong CM, Edelman GM. Expression of cell-adhesion molecules in embryonic induction. I. Morphogenesis of nestling feathers. J Cell Biol 1985; 101:1009-26. 58. Chuong CM, Edelman GM. Expression of cell-adhesion molecules in embryonic induction. II. Morphogenesis of adult feathers. J Cell Biol 1985; 101:1027-43. 59. Itow T. Inhibitors of DNA synthesis change the differentiation of body segments and increase the segment number in horseshoe crab embryos. Roux’s Arch Dev Biol 1986; 195:323-33. 60. Patel NH, Kornberg TB, Goodman CS. Expression of engrailed during segmentation in grasshopper and crayfish. Development 1989; 107:201-12. 61. Irvine KD, Wieschaus E. Cell intercalation during Drosophila germband extension and its regulation by pair-rule segmentation genes. Development 1994; 120:827-41. 62. Boissonade J, Dulos E, DeKepper P. Turing patterns: From myth to reality. In: Kapral R, Showalter K, editors. Chemical Waves and Patterns. Boston: Kluwer, 1994:221-68. 63. Harding K, Hoey T, Warrior R et al. Autoregulatory and gap gene response elements of the even-skipped promoter of Drosophila. EMBO J 1989; 8:1205-12. 64. Ish-Horowicz D, Pinchin SM, Ingham PW et al. Autocatalytic ftz activation and instability induced by ectopic ftz expression. Cell 1989; 57:223-32. 65. Goto T, MacDonald P, Maniatis T. Early and late periodic patterns of even skipped expression are controlled by distinct regulatory elements that respond to different spatial cues. Cell 1989; 57:413-22. 66. Stanojevic D, Small S, Levine M. Regulation of a segmentation stripe by overlapping activators and repressors in the Drosophila embryo. Science 1991; 254:1385-7. 67. Guthrie S, Lumsden A. Formation and regeneration of rhombomere boundaries in the developing chick hindbrain. Development 1991; 112:221-9.

358

Molecular Basis of Epithelial Appendage Morphogenesis

68. Guthrie S, Prince V, Lumsden A. Selective dispersal of avian rhombomere cells in orthotopic and heterotopic grafts. Development 1993; 118:527-38. 69. Heyman I, Kent A, Lumsden A. Cellular morphology and extracellular space at rhombomere boundaries in the chick embryo hindbrain. Dev Dyn 1993; 198:241-53. 70. Gierer A. Physical aspects of tissue evagination and biological form. Quart Rev Biophys 1977; 10:529-93. 71. Mittenthal JE, Mazo RM. A model for shape generation by strain and cell-cell adhesion in the epithelium of an arthropod leg segment. J Theoret Biol 1983; 100:443-83. 72. Fristrom D, Chihara C. The mechanism of evagination of imaginal discs of Drosophila melanogaster. V. Evagination of disc fragments. Dev Biol 1978; 66:564-70. 73. Maniotis AJ, Chen CS, Ingber DE. Demonstration of mechanical connections between integrins, cytoskeletal filaments, and nucleoplasm that stabilize nuclear structure. Proc Nat Acad Sci USA 1997; 94:849-54. 74. Yurchenco PD, O’Rear JJ. Basal lamina assembly. Curr Opin Cell Biol 1994; 6:674-81. 75. Waddington CH. The Strategy of the Genes. London: Allen and Unwin, 1957. 76. Newman SA. Interplay of genetics and physical processes of tissue morphogenesis in development and evolution: The biological fifth dimension. In: Beysens D, Forgacs G, Gaill F, editors. Interplay of Genetic and Physical Processes in the Development of Biological Form. Singapore: World Scientific, 1995:3-12. 77. Meinhardt H. Cell determination boundaries as organizing regions for secondary embryonic fields. Dev Biol 1983; 96:375-85. 78. Newman SA. Sticky fingers: Hox genes and cell adhesion in vertebrate limb development. BioEssays 1996; 18:171-4. 79. Chuong CM. The making of a feather: homeoproteins, retinoids and adhesion molecules. BioEssays 1993; 15:513-21.

CHAPTER 18

Periodic Pattern Formation of the Feathers Han-Sung Jung, Cheng-Ming Chuong

Introduction

A

major question in embryonic development is how cells and tissues become precisely arranged to make up the body plan. It is best to study this process in an organ with well defined morphological patterns. One of the simplest patterns observed is the maintenance of a minimum distance between neighboring elements, which is often called a “spacing pattern”.1,2 In a spacing pattern, some cells from a field of originally equivalent precursor cells become different from their neighbors, thus forming a spaced array of determined cells. Examples of spacing patterns are found in the development of many vertebrate organs, as in somite formation,3,4 tooth formation,5-7 skin gland patterning,8 scale formation9 and alligator skin patterning.10 Related to such spacing patterns are slightly more complex ones where the pattern is anisotropic, which might be included in a more sophisticated pattern than the model of isotropy, such as the stripes seen in zebra and in fish.11 A favored model for pattern formation is avian feather buds that are arranged in a highly ordered array.12 In the following we will use this model for discussion (Figs. 18.1, 18.2). A more detailed description of feather development is described in chapter 5. Briefly, the skin of the bird is divided into tracts, the feathered regions and apteria, the naked regions (Fig. 18.1). Within the tracts, there are initially no morphological nor histochemical indicators of periodic patterns. For the spinal tract, the first three or four primordia then appear along the midline in the posterior lumbar region at stage 29.13,14 This row first extends in both directions by the addition of successive, closely spaced primordia. In the anterior lumbar region, where the medial area remains bare, two initial rows of primordia form, one on either side of the midline. Later, successive rows are added parallel and lateral to these initial rows at approximately six hour intervals. Each new primordium lies between, and close to, two neighbors in the preceding row. The pterylae have similar temporal and spatial patterns of development, as first described by Holmes (1935),15 and later studied in more detail for the humeral, femoral and spinal tracts.16 For each feather primordium, both the dermis and epidermis play essential roles. Which one acts first? Morphologically, the epidermal placode is observed earlier than the dermal condensation in the midline of the back skin.17 However, this may be biased by our inability to observe earlier changes in the dermis, and there is more evidence suggesting that dermis plays the initial role (see chapter 5 for further discussion).

Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

360

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 18.1. Overview of feather patterns in the dorsal skin of 10 day old chick embryo. This embryo shows: (1) the characteristic distribution of different tracts, or pterylae, on the skin, and (2) the regular periodical arrangement of feather primordia in each tract. Ap, apterium; AT, alar tract; CT, caudal tract; FT, femoral tract; HT, humeral tract; mAp, middorsal apterium; ST, spinal tract.

The regular spacing of feather patterns has intrigued developmental biologists and mathematical biologists for decades. An explanation of the mechanism of feather pattern formation should take into account the following phenomena: 1. the periodic nature of the pattern; 2. the sequential formation of the pattern; 3. the position of new individual primordia; and 4. the inter-primordial spacing. In this chapter, we will first have an overview of previous models which include lateral inhibition, maximal space filling, reaction-diffusion mechanism, mechano-chemical model, etc. Then, based on our recent molecular findings, we will present a model that also draws from these previous perspectives.

Surveys of Existing Models Lateral inhibition is the process through which one developing element prohibits the development of similar elements nearby. This was first suggested by Wigglesworth (1940)19 for the regular spacing of bristles in the bug Rhodnius. Here, each bristle site, as it formed, utilized precursors from the surrounding tissue and, by competition, inhibited bristle formation within a certain surrounding area. Later, well-characterized studies on the patterning of sensory bristles in the epidermis of Drosophila were carried out.20 The molecular

Periodic Pattern Formation of the Feathers

361

Fig. 18.2. Schematic drawing of 9 day old chick embryo. (A) Dorsal view of nine day old chick dorsal pteryla showing complete pattern of feather buds. The initial feather buds are drawn in solid black. Note that there are two initial rows in the cervical and thoracic regions, but one mid-dorsal initial row in the lumbar, sacral and caudal regions. Ant: Anterior; Post: Posterior. (B) Illustration of early developmental stages of feather buds. A, First noticeable changes in feather-forming region convert the ectoderm into columnar epidermis and loose mesenchyme into dense dermis. B, Epidermis then thickens into a placode and underlying dermis becomes even more dense. After induction occurs between ectoderm and mesenchyme, the two components become epidermal placode and dermal condensation respectively. C, Feather bud has an anterior-posterior (A-P) polarity, and this A-P axis is referred to as the ‘Feather axis’. D, Elongated feather bud. Prox: Proximal; Dist: Distal.

basis of spacing patterns is better studied in Drosophila because of the availability of mutants. Recent molecular studies led to the suggestion that the activity of the sevenless tyrosine kinase pathway may be involved in the determination of the spacing pattern of ommatidia in the Drosophila eye.21,22 Although exactly how precluster cell groups come to form in a regular spatial array is unknown, lateral inhibition by Scabrous is likely to be involved. For example, recent data suggest that Scabrous is involved in this process by preventing an excess of cells from adopting a neural precursor fate.23 Another well characterized molecular mechanism producing lateral inhibition is the Notch pathway used in Drosophila sensory bristle pattern formation and many other systems.24,25 It is believed that a cell with potential to become a neuron expresses Delta, which binds to its receptor, Notch, on neighboring cells. This triggers a response, effectively prohibiting these cells from becoming neurons. In the mean time, the synthesis of Delta in these cells is suppressed so that their ability to deliver inhibition through Delta is diminished. This gives rise to a feedback loop that can

362

Molecular Basis of Epithelial Appendage Morphogenesis

amplify the unstable temporal or spatial differences between adjacent cells and set up the evenly spaced sensory bristles. In the above insect based cases, the interactions occur within one single layer of epithelium. In vertebrates, the skin appendages involve both epithelium and mesenchyme, as well as multiple levels of tissue interactions. Using a similar scheme to study the pattern of wool follicles in sheep skin, Claxton (1973)26 suggested that if one takes account of skin growth and the possible variation in the radius of effective inhibition, it is possible to simulate the patterning process. Realistic values for the range and variance of nearest neighbor distances within the pattern can be achieved. In the same line, Ede (1972)27 proposed a model for feather pattern formation. According to this model, a zone of inhibition is generated radially around each forming feather primordium, by the production of hypothetical diffusible chemicals that inhibit feather formation. New primordia would only form outside the circular threshold contour of inhibition around existing sites. If inhibition decreases steeply beyond the threshold contour, and some mechanism exists to generate primordia outside the zone of inhibition, these conditions together would give the pattern a well-defined periodicity. To account for the sequential appearance of feather primordia, Ede (1972)27 suggested that the competence to form primordia is confined to a narrow, but ever-widening, band of dermis lateral to already-formed primordia. So, new primordia would form as soon as sufficient competent dense dermis became available. Ede suggested that the spread of competent dermis may depend on the lateral diffusion of an activator substance from the midline. However, this hypothetical long range action would result in an exponential decrease in the rate of initiation of new rows, with increasing distance from the midline. In fact, initiation appears to be approximately constant, at least over the first few rows. This difficulty could be overcome by suggesting that a key substance is produced by successive bands of activated dermis that act with a short range, or by the development of competence across the tract immediately in advance of morphogenesis. Sengel (1976)18 interprets this as a “wave of morphogenetic activity”. Typically, it starts from the primary row and spreads through the tracts. However, there is also evidence that lateral rows can form in cultured explants that do not contain the primary row.28 A piece of thigh skin from before the onset of morphogenesis was cultured. A normal spacing pattern developed in this piece of skin; the first row formed at the margin which had been nearest the presumptive site of the primary row in the intact tract. Furthermore, experiments in epithelial-mesenchymal recombination were carried out on skin explants from stage 29-34 chick embryos. The location is always under dermal control.28,29 When the midline containing the primary row is removed and the epithelia and mesenchyme are recombined, dermal condensations at this stage are unstable and a new primary row emerges from the region where the morphogenetic wave is located before recombination.29 What happens within this “morphogenetic wave”? Sengel (1976)18 proposed that feather buds are patterned in a hexagonal array because this arrangement accommodates a maximal number of elements in a given area. The models therefore set out to explain the pattern using space-filling constraints. The geometric regularity of this approach and the assumption that primordia are equivalent imply that each new primordium is equidistant from two neighbors in the preceding row. This is best demonstrated in the en face sections of primordia forming dermis. Histological sections indicate that, between formed primordia, elongated dermal cells are aligned in parallel to form “arrays”.13,30 These arrays intersect at sites of dermal condensations. Extracellular collagen fibrils are also aligned along these arrays. Cells may move along these fibrils in these “cellular highways” to intersections where their movement is restricted and there they form clusters which enlarge into condensations. The arrangement of extracellular matrix and dermal cells becomes progressively organized in

Periodic Pattern Formation of the Feathers

363

the dermis lateral to the already formed condensations, so that diagono-lateral arrays intersect at the sites of new primordia. Sequential appearance is then accounted for by the progressive organization of the dermis into arrays which extend diagono-laterally from formed condensations to generate a lattice of oriented cells and fibrils in hitherto unpatterned skin. The intersections in this lattice define the sites of new primordia. In this template-like mechanism, primordia will be laid out in order once the lateral propagation starts. However, these models do not suggest how the periodic structure of the primary row forms, nor do they point out how the patterning is initiated. Additionally, in some cases the primordia of skin appendages do not form in close approximation, such as in lamb’s wool, reptilian scales and avian scales.9,26,31 There is no histological evidence that dermal structures are preformed through direct interactions between primordia. A different mechanism of pattern determination may operate in these situations. Periodic patterns can also be generated by the differential diffusion of chemical substances. Turing32 showed how an initially homogeneous system of two or more diffusible chemical “morphogens” could develop periodic heterogeneity after small, random disturbances. He then suggested that such a chemical distribution could form the basis of periodic patterns. The new concept also gave rise to the idea that diffusible signaling molecules in combination with random intrinsic instability may be enough to generate spacing patterns in a biological system. Meinhardt33,34 further suggested that randomly generated initiation sites can produce both diffusible activators and inhibitors. With activators acting within a short range and inhibitors acting at a long range, it is possible to generate a stable periodic pattern. Nagorcka and co-workers also have applied these principles to explain the arrangement of mammalian hair follicles.35 Later, it was proposed that the Turing model does not have to be limited to chemical substances. It is also possible to generate spacing patterns by mechano-chemical force behaving in a Turing fashion.36,37 From a homogeneous dense dermis, groups of mesenchymal cells break up into a row of dermal condensations. As feather buds form, tension lines develop, joining centers of dermal condensations. This change leads to spatial instability which generates new feather buds. The change is based on the properties of dermal mesenchymal cells in vitro. These cells spread and migrate within an environment consisting of fibrous extracellular matrix and other cells. They respond to haptotaxes which direct cell movements in response to adhesion gradients on their substratum or neighboring cell membranes. As these cells move, they generate large traction forces that further organize the elastic extracellular matrix and facilitate future cell movement in particular spatial patterns. Depending on the anisotropic character of the substratum or cell adhesive properties, the final periodic array can exhibit a variety of geometric patterns. Many of the models presented above are based on phenomenological data, and the molecular identities of the parameters in these models are largely unknown. Later, the importance of adhesion molecules was demonstrated. It was found that the extracellular matrix is enriched for collagens with specific distributions of fibronectin, proteoglycan and tenascin.38-40 NCAM is initially also found to be uniformly distributed in the dense dermis, and later enriched in the dermal condensation but absent in the inter-primordial dermis.41 Antibody perturbation of NCAM, tenascin and integrin show that they are involved in different stages of the dermal condensation process.42,43 Similarly, inhibitors of proteoglycan and collagen processing led to abnormal feather bud morphogenesis.44,45 These results show that these adhesion molecules are involved. Still, exactly how they work to initiate, propagate and stop an individual dermal condensation and to arrange themselves into patterns remains to be learned. Recently, many signaling molecules have been found in different developing models and their effects in feather morphogenesis are discussed in chapter 13.

364

Molecular Basis of Epithelial Appendage Morphogenesis

Here we will use the available information to present a working model with some molecular candidates.

A Novel Integrated Model of Feather Pattern Formation We and others have been studying the roles of some of these key molecules, namely FGF, Shh, BMPs, etc. in feather pattern formation.46-48 Based on these data and ideas from the previous models, we will present a model that includes a computer simulation (Fig. 18.3). The summary of these data is listed here to provide the rationale of our model: 1. Shh and FGF-4 favor the formation of feather domains in skin explant cultures. 2. BMP-2 and BMP-4 suppress formation of the feather domain in explant cultures. 3. The expression of Shh and FGF start as a continuous linear pattern in the midline, progressing from the posterior to the anterior, and then changing to a punctate pattern. 4. BMP-2 and BMP-4 start to be expressed a little bit later, directly in the feather bud domain. The goal is to have a model that can satisfy the following three aspects of skin appendage patterning in the lumbar region where we have experimental data. First, the first feather

Fig. 18.3 (opposite). Model depicting several steps along the feather patterning cascade. In this model, feather patterning is regulated by: (1) a wider distribution of activator at the tract level, (2) a gradient of competence to form skin appendages that advances from posterior to anterior (in the case of lumbosacral region of the spinal pterylae, and (3) local lateral inhibition triggered by the putative feather primordia that sets up interbud spacing and leads to individual feather primordia in a periodic fashion within the competent field. (A-D) Top view. (A) During development, the gradient of competence (elliptical curve) traverses the skin in a posterior to anterior direction before feather buds form. Where the competence signal meets the midline stripe, a feather initiation site is formed (marked by X). Although the existence of this competence wave is consistent, its molecular nature is yet to be identified. Since the ligands of activators already exist in the midline stripe, we hypothesize this competence to be the competence to respond to the activators (e.g., expression or conformation changes of growth factor receptors, or signaling molecules downstream to growth factors). (B) From the feather initiation site, both activators (small circle/dot) and inhibitors (large circle) are released locally and diffuse into the surrounding regions. BMPs may be considered long range morphogens and Shh a short range morphogen,51,52 while the diffusion of FGF may be slowed down by binding to extracellular matrix.53 Therefore, in this model, the inhibitors are likely to be distributed more widely than the activators. (C) As the gradient of competence continues to travel anteriorly, and as soon as the competence signal passes outside of the inhibitory field, another feather initiation site is formed (X). Processes (B) and (C) then repeat. This leads to the propagation of feather buds and the conversion from the linear to periodic pattern, thus forming the primary row. (D) As the gradient of competence moves along the midline, it also spreads bilaterally. Therefore, the competence signal moves in a generally half elliptical curve. Processes similar to (A, B, and C) repeat in the medial-lateral fashion, thus forming the secondary rows and further rows. (E) Developmental sequences viewed from the lateral upper side to summarize the model. The gradient of competence moves over time (t1, t2 and t3). When the feather bud initiation tendency (activation of the activator signaling pathway) is above the threshold, a presumptive feather primordium is initiated and becomes a discrete feather bud through a local positive regulatory loop. The simultaneously secreted inhibitors lead to the formation of the interbud space. A computer program based on these principles was prepared in which simulated feather bud formation occurs sequentially postero-anteriorly as well as bilaterally.

Periodic Pattern Formation of the Feathers

365

A

B

Ant

Post Lateral

Midline

Lateral

D

C

E Lat

Threshold

t1 Post

t2

t3 Ant Midline

Lat

366

Molecular Basis of Epithelial Appendage Morphogenesis

bud is initiated posteriorly in the dorsal midline of the lumbo-sacral region. Second, more feather buds are added anterior to the first one sequentially along the midline with regular spacing. Third, feather buds in lateral rows are formed in between two feather buds of the midline row. With experimental data, we propose that FGF and Shh are candidate activators, while BMP-2 and BMP-4 are candidate inhibitors. Sequential formation of feather buds is preceded by the orderly expression of signaling molecules. We report the following observations: 1. There are physiological diffusible chemicals that work as activators and inhibitors. 2. Both activators and inhibitors diffuse out from the same source, in favor of a reaction-diffusion mechanism and lateral inhibition. 3. Activators appear before inhibitors. 4. Activators appear in a linear pattern and then assume a periodic distribution. 5. Inhibitors appear directly in a periodic pattern. The following model is built upon a reaction-diffusion mechanism, lateral inhibition, and the propagation of a competence gradient which progresses with time. The feather tract is initiated with global activators expressed as a continuous stripe in the midline (Fig. 18.3A-D). Within the feather tract field, there is a position-dependent gradient specifying competence to form a feather bud. For the lumbo-sacral region of the spinal tract, the gradient has its peak at the posterior end of the midline. We hypothesize that competence is progressively gained (under the control of a mechanism yet to be identified, but which may originate from the Hox gradient in the skin).49 A series of steps may then occur. As soon as the competence gradient allows cells to respond to the global activators (which already exist), the first feather bud is initiated, resulting in the local synthesis of activators and inhibitors which diffuse into the surrounding regions. The activators have a higher potency but shorter range of action than the inhibitors, while the inhibitors diffuse further than the activators and have a longer range of action. The end result is an activation zone immediately adjacent to the initiation sites with an inhibitory zone outside it, thus setting up the spacing pattern. Since there is a gradient of competence which has its peak at the posterior end, the second feather bud is initiated anterior to the first one in the midline. There is also the hypothetical expansion of the competence gradient from the midline toward the lateral edge. Feather buds are initiated lateral to the primary row in the nearest place lateral to the feather buds in the midline. A diagram of developmental sequences viewed from the lateral side to summarize the model is shown in Figure 18.3 E. The competence field moves over time (t1, t2, t3...). When the feather initiation potential triggering the activator signaling pathway is above the threshold, a presumptive feather site is initiated. The simultaneously secreted inhibitors lead to the formation of the interbud space. The relative amount of activators and inhibitors set up the boundary of a feather primordial domain, which then becomes a discrete feather bud. The gradient that controls the sequential development of competence to form feather buds along the midline may be set early in development in relation to positional information gradients as proposed by Wolpert.50

Conclusions To summarize, periodic pattern formation is a fascinating phenomenon, particularly obvious in the formation of epithelial appendages. Feather pattern formation provides a model that can be tested with experimental embryology and molecular biology approaches. The strength of the model we present is that we have incorporated the most recent experimental data to offer some molecular candidates for some of the parameters. However, the model is far from completion. Like previous models, some hypothetical parameters have to be used to build the model. However, we are glad to make an initial effort to integrate the

Periodic Pattern Formation of the Feathers

367

experimental data and spacing models. We still need more experimental data to revise and improve this working model and we hope the principles used here for feather patterning will serve as an example for more understanding of integument patterning.

Acknowledgment Part of this work is H-S.J.’s Ph.D. thesis with Professor Lewis Wolpert. We would like to express our gratitude to Dr. Wolpert. We also thank Drs. Randall B. Widelitz and C. Tickle for helpful discussion. The work is partially supported by grants from NIH and NSF.

References 1. Wolpert L. Positional information and pattern formation. Curr Top Dev Biol 1971; 6:183-224. 2. Wolpert L, Stein WD. Positional information and pattern formation. In Pattern Formation: A Primer in Developmental Biology. Maklacinski GM, Bryant SV eds. Macmillan, 1984:3-21. 3. Pearson M, Elsdale T. Somitogenesis in amphibia. II. Origins in early embryogenesis of two factors involved in somite specification. J Embryol Exp Morphol 1979; 53:245-267. 4. Davidson D. Segmentation of frogs. Dev Supp 1988; 104:221-229. 5. Osborn JW. The ontogeny of tooth succession in Lacerta vivipara Jacquin (1787) Proc. R. Soc. Lond. B. 1971; 179:261-289. 6. Osborn JW. Morphogenetic gradients: Field versus clones. In: Butlerm PM, Joysey KA eds. Development, function and evolution of teeth. London: Academic Press, 1978:171-201. 7. Smith MM. Evolutionary change and heterochrony of enemel in vertebrates. In Heterochrony in the Evolution. Mcnamara KJ ed. John Wiley and Sons Ltd, 1995:125-150. 8. Holder N, Glade R. Skin glands in the axolotol: the creation and maintenance of a spacing pattern. J Embryo Exp Morph 1984; 79:97-112. 9. Maderson PFA. The embryonic development of the squamate integument. Acta. Zoologica 1965; XLVI: 275-295. 10. Murray JD. Mathematical Biology. 2nd edition. Berlin: Springer-Verlag, 1993. 11. Kondo S, Asai R. A reaction-diffusion wave on the skin of the marine angelfish Pomacanthus. Nature 1995; 376:765-768. 12. Sengel P. Pattern formation in skin development. Int J Dev Biol 1990; 34: 33-50. 13. Stuart ES, Garber B, Moscona AA. An analysis of feather germ formation in the embryo and in vitro, in normal development and in skin treated with hydrocortisone. J Exp Zool 1972; 179:97-118. 14. Davidson D. The mechansim of feather pattern development in the chick. I. The time determination of feather position. J Embryol Exp Morph 1983; 74:245-259. 15. Holmes A. The pattern and symmetry of adult plumage units in relation to the order and locus of orign of embryonic papillae. Am J Anat 1935; 56:513-535. 16. Mayerson PL, Fallon JF. The spatial pattern sequnece in which feather germs arise in the white Leghorn chick embryo. Dev Biol 1985; 109:259-267. 17. Sengel P. Feather pattern development. Ciba Foundation Symposium, No 29. Elsevier, 1978. 18. Sengel P. Morphogenesis of Skin. Cambridge: Cambridge University Press, 1976. 19. Wigglesworth VB. Local and general factors in the developmental of pattern in Rhodnius Prolixus (hemiptera). J Exp Biol 1940; 17:180-200. 20. Lawrence, P. The Making of a Fly. London: Blackwell Scientific, 1992. 21. Rubin GM. Signal transduction and the fate of the R7 photoreceptor in Drosophila. Trends in Genetics 1991; 7:372-377. 22. Hafen E. Patterning by cell recruitment in the Drosophila eye. Currrent Opinions in Genetics and Development 1991; 1:268-274. 23. Ellis MC, Weber U, Wiersdroff V, Mlodzik M. Confrontation of scabrous expressing and non-expressing cells is essential for normal ommatidial spacing in the Drosophila eye. Development 1994; 120:1959-1969.

368

Molecular Basis of Epithelial Appendage Morphogenesis

24. Collier RC, Monk NAM, Maini PK,Lewis JH. Pattern formation by lateral inhibition with feedback: A mathematical model of Delta-Notch intercellular signalling. J Theo Biol 1996; 183:429-446. 25. Artavanis-Tsakonas S, Matsuno K, Fortini ME. Notch signaling. Science 1995; 268:225-268. 26. Claxton JH.A model of pattern formation in the primary skin follicle population of sheep. J Theor Biol 1973; 40:353-367. 27. Ede DA. Cell behaviour and embryonic development. Int J Neurosci 1972; 3:165-174. 28. Linsenmayer T. Control of integumentary patterns in the chick. Dev Biol 1972; 27:247-271. 29. Novel G. Feather pattern stability and reorganization in cultured skin. J Embryol Exp Morph 1973; 30:605-633. 30. Stuart ES, Moscona AA. Embryonic morphogenesis; role of fibrous lattice in the development of feather and feather patterns. Science 1967;157:947-948. 31. Sawyer RH. Avian scale development: I. Histogenesis and morphogenesis of epidermis and dermis during formation of the scale ridge. J Exp Zool 1972; 181:365-384. 32. Turing A. The chemical basis of morphogenesis. Phil Trans Roy Soc London. 1952; B237:37-72. 33. Meinhardt H. (1982). Models for biological pattern formation. London: Academic Press, 1982. 34. Koch AJ, Meinhardt H. Biological pattern formation: From basic mechanisms to complex structures. Rev Mod Phys 1994; 66:1481-1508. 35. Nagorcka BN, Mooney JR. The role of a reaction-diffusion system in the initiation of primary hair follicles. Theor Biol 1985; 114:243-272. 36. Murray JD, Oster GF, Harris AK. A mechanical model for mesenchymal morphogenesis. J Math Biol 1983; 17:125-129. 37. Oster GF, Murray JD, Harris AK. Mechanical aspects of mesenchymal morphogenesis. J Embryol Exp Morph 1983;78:83-125. 38. Mauger A, Demarchez M, Herbage D et al. Immunofluorescent localization of collagen type I and III, and firbonectin during feather morphogenesis in the chick embryo. Dev Biol 1982; 94:93-105. 39. Kitamura K. The structure and distribution of chondrotin sulphate during the formation of the chick embryo feather germ. Development 1987; 100:501-512. 40. Tucker RP. The sequential expression of tenascin mRNA in epithelium and mesenchyme during feather morphogenesis. Roux Arch Dev Biol 1991; 200:108-112. 41. Chuong CM, Edelman GM. Expression of cell adhesion molecules in embryonic induction. I. Morphogenesis of nestling feathers. J Cell Biol 1985; 110:1009-1026. 42. Gallin WJ, Chuong CM, Finkel LH, Edelman GM. Antibodies to liver cell adhesion molecule perturb inductive interactions and alter feather pattern and structure. Proc Natl Acad Sci 1986; 83:8235-8239. 43. Jiang TX, Chuong CM. Mechanism of feather morphogenesis: I. Analyses with antibodies to adhesion molecules tenascin, N- CAM and integrin. Dev Biol 1992; 150:82-98. 44. Goetinck PF, Carlone DL. Altered proteoglycan synthesis disrupts feather pattern formation in chick embryonic skin. Dev Biol 1988; 127:179-186. 45. Marsh RG, Gallin WJ. Toxic effects of beta-aminopropionitrile treatment on developing chicken skin. J Exp Zool 1994; 268:381-389. 46. Widelitz RB, Jiang TM, Noveen A et al. FGF induces skin appendages from developing skin. J Invest Dermatol 1996; 107:797-803. 47. Song HK, Wang Y, Goetinck PF. Fibroblast growth factor-2 can replace ectodermal signaling for feather development. Proc Natl Acad Sci USA 1996; 93:10246-10249. 48. Ting-Berreth S, Chuong CM. Sonic hedgehog in feather morphogenesis: Induction of mesenchymal condensation and association with cell death. Dev Dyn 1996; 207:157-170. 49. Chuong CM, Oliver G, Ting S, Jegalian B, Chen HM, De Robertis EM. Gradient of homeoproteins in developing feather buds. Development 1990; 110:1021-1030. 50. Wolpert L. Positional information and the spatial pattern of cellular differentiation. J Theore Biol 1969; 25:1-47.

Periodic Pattern Formation of the Feathers

369

51. Lawrence PA, Struhl G. Morphogens, compartments, and pattern: Lessons from drosophila? Cell 1996; 85:951-961. 52. Lecuit T, Brook WJ, Ng M et al. Two distinct mechanisms for long-range patterning by Decapentaplegic in the Drosophila wing. Nature 1996; 381:387-393. 53. Aviezer D, Hecht D, Safran M et al. Perlecan, basal lamina proteoglycan, promotes basic fibroblast growth factor-receptor binding, mitogenesis, and angiogenesis. Cell 1994; 79:1005-1013. 54. Jung et al. Local inhibitory action of BMPs and inter-feather bud spacing: A model for periodic pattern formation. Dev Biol 1998; 196:11-23.

CHAPTER 19

Gene Networks and Supernetworks: Evolutionarily Conserved Gene Interactions Alexander Noveen, Volker Hartenstein, Cheng-Ming Chuong

Introduction

M

any genes involved in development are evolutionarily conserved and interact with other genes either directly or indirectly. More recently it has become clear that gene interactions are also evolutionarily conserved. Here a model of gene interaction is presented. Many interacting genes give rise to a gene network. Two gene networks are homologous (i.e., have the same ancestry) if they have genes which are homologous and interact with each other in similar ways. Many interacting gene networks can give rise to a gene supernetwork. The function of a gene supernetwork is more complicated than a gene network. A gene supernetwork, for example, may be involved in determining the development of an entire limb while a gene network, working within the supernetwork, may be involved in setting up one of the axes of the limb bud (i.e., anterior-posterior, dorsal-ventral, and proximal-distal). Developmental programs present today in various species are a product of millions of years of evolution. Species have to possess the ability to change such programs as a result of the demands of new environments. The ability to generate phenotypic diversities will ensure the adaptive success of a species. Much evidence indicates that during evolution, diversities in developmental programs are usually derived from variations in old programs. One of the most dramatic examples of evolution of phenotypic diversity is the presence of various integument appendages in vertebrates. As described in the introductory chapter 1, these appendages range from scales, feathers and hairs to horns, teeth and glands. Such appendages all begin as an epithelial placode that grows out and forms an epithelial appendage. These epithelial appendages appear to develop as a result of the activity of very similar genes. A question then that arises is: Why are the shapes of these appendages so different if they rely on very similar genes? The answer, as will be discussed later, seems to be that, although the same genes are used during the development of different epithelial appendages, gene expressions (both spatial and temporal) and interactions are different. Since the advent of protein and DNA sequencing techniques, much effort has been devoted to comparison of gene sequences. Such comparisons have revealed that most genes have sequences that are evolutionarily conserved and can be found in other related genes. Based on their sequence homologies, related genes can be assigned to a specific gene family. Here we will present a model and review evidence that suggests that, similarly to their conserved sequences, genes have interactions with each other that are also evolutionarily conserved. Based on their conserved interactions, networks of genes can be assigned to specific gene network families. Gene networks within each family are descended from the same

Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

Molecular Basis of Epithelial Appendage Morphogenesis

372

ancestral gene network (Fig. 19.1) and can be distinguished from other families based on their sequences and interactions. A gene network usually carries out developmental tasks in association with other gene networks. Such an assembly of gene networks is called a gene supernetwork. Gene supernetworks are responsible for the formation of an entire organ, segment, limb or skin appendage. Although many processes such as metabolism, immune response and cell cycle are regulated by conserved gene networks and supernetworks, here we will focus on those that are specifically involved in development. In explaining the gene network and supernetwork models we will emphasize similarities in developmental programs among species as well as similarities in development of various structures in the same species. We will first review some findings about gene evolution on which gene network and supernetwork models are based.

Genes Conservation of Gene Structure and Function It appears that a large portion of presently existing genes are descendants of genes that evolved before the divergence of vertebrates and invertebrates. There also exists an abundance of genes that evolved before the divergence of prokaryotes and eukaryotes, pointing to the ancient origins of such genes.1 About 40-50% of the proteins that have been sequenced so far contain segments which have homologies with other proteins and thus can be placed into specific protein families.2 The rest of the sequenced proteins apparently have no homologies with other proteins. A portion of these proteins will undoubtedly be found to belong to specific protein families as new genes are sequenced. The number of extant protein families has been estimated to be about a thousand. This estimate has been reached based on comparing primary structure3-8 and tertiary structure9,10 of proteins. In the future, tertiary structure comparisons will probably help to identify more related proteins that have little primary structure similarity. While there is a possibility that the actual number of unique protein families is a little more or less than the estimated number, the fact remains that there is a limited number of protein families found in nature. The proteins that show no sequence similarity with other proteins either are evolutionarily unique or descended from older proteins, but have gone through many mutations and lost any similarity to their ancestors. Among the above two alternatives, the second is more favored.1,11 It is speculated that very early in the history of gene evolution, there were a limited number of protein domains that had evolved specific functions. Once such domains were present, evolution of new protein domains became minimal because new proteins (with new functions) could be generated from an established reservoir of already existing proteins and protein domains.1,7,11 Thus, the present day proteins that have no similarity to other proteins are probably related to the extant protein families but have lost any recognizable structural similarity. Conservation of gene structure and function usually go hand in hand. The conservation of the structure of a gene usually means that the function of the gene is also conserved.12 However, it should be noted that in the same way that new proteins evolve from old proteins, new functions also evolve from old functions. Thus a protein with a specific function could be recruited to perform another function (For example, see refs. 13,14).

Evolution of New Genes There are three major ways that new genes can evolve.

Gene Networks and Supernetworks

373

Fig. 19.1. The hypothetical evolution of a gene network family. Through duplication of an ancestral gene network and further structural changes in gene members (such as gain of function mutations), new gene networks can form. Each of the geometric figures represents a different gene. Figures that look similar are homologous (having the same ancestry) to each other. The sharp and blunt arrows indicate that the upstream gene is respectively stimulating or inhibiting the activity, availability, expression, or translation of the downstream gene through direct or indirect protein interactions. Downstream genes may also regulate the activity or expression of upstream genes, as is shown here in gene network 4 and the ancestral gene network. This is a very simplified model. For example, here each gene network is shown to be composed of 3-6 members. In reality, there may be approximately 50-200 genes participating in any gene network. Differences due to mutations and other changes in each gene are shown by different fill-ins. The four resulting gene networks are found in the same species and are thus paralogous (having homology in the same species) to each other. They can be identified as paralogous because their gene members are paralogous to each other and have the same interactional circuitry (i.e., the ways they interact with each other are similar). However, during evolution both homology and circuit changes may occur as shown for the resulting four gene networks. Note that only gene network 4 has retained its ancestral circuitry. Structural changes (including mutations, additions, and deletions) have occurred for all the genes. Also note that gene networks 1, 2, and 3 have each lost three of their gene members, while gene network 3 has gained two new members. Gene networks 1 and 2 have closer ancestry than networks 3 and 4. These networks may have similar or different functions, depending on the time since their duplication and divergence. During evolution, other changes, i.e., association of two networks which have different ancestry, may occur (not shown).

374

Molecular Basis of Epithelial Appendage Morphogenesis

Gene duplication and divergence A new copy of an already existing gene can appear in the genome by duplication of a segment of DNA that contains that gene.15 Duplication of a DNA segment can occur frequently, mostly due to unequal crossover. Gene duplication usually refers to the duplication of both the coding and regulatory sequences of a gene. During their evolution, duplicated genes may structurally diverge and their ancestral function may be modified. Since they share the same ancestor, they are homologous to each other. In some cases, due to structural defects in their regulatory or coding regions, duplicated genes may become inactive and turn into pseudogenes. In the same species, homologous genes that are produced as a result of gene duplication are called paralogs. Homologous genes in two different species, having the closest sequence similarity and function, are called orthologs and are a result of speciation. A gene in one species that has similar DNA sequences to another gene in another species may not necessarily be the ortholog of that gene. Thus the assignment of orthology may sometimes require close examination of other family members. Duplication of genes can also occur by polyploidization of chromosomes. Comparisons of vertebrates with invertebrates like Drosophila melanogaster and Caenorhabditis elegans indicate that vertebrates usually have four times as many genes as invertebrates.16 Additionally, vertebrates usually have gene families with four members, while invertebrates have only one ortholog.17 This indicates that vertebrates have undergone two rounds of genome duplication since their divergence from invertebrates.17-19 Although in many cases they have lost one or more of the duplicated copies (for example vertebrate hedgehog and Msx families have each lost one member), in some cases they have retained all of them. An outstanding example is Hox genes, which are found in four paralogous clusters.20 On the other hand, some gene families like Wnt,21 bone morphogenetic protein (BMP)22 and fibroblast growth factor (FGF)23 have more than four members. These gene families were probably produced as a result of both individual gene duplication and polyploidization. Exon shuffling Some exons code for protein domains that can move from one gene to another.24,25 A protein domain or fold is defined as a part of a protein that can form a tertiary protein structure independent of its neighboring amino acid sequences. Protein domains that are evolutionarily mobile are called modules.26 In more recent stages of protein evolution, during the evolution of higher eukaryotes, some of the newly evolved proteins have been formed from a combination of modules.27 For example, mosaic proteins such as cell adhesion molecules, extracellular matrix proteins, proteins in body fluids and cell surface receptors are made up of two or more modules.7 It should be noted, however, that most exons do not code for a whole domain. Usually protein domains are coded by two or more exons separated by introns. Thus in order to keep the integrity of a module, more than one exon (and their introns) must be shuffled. Although it appears that during the metazoan evolution, introns have been greatly involved in exon shuffling, intronic recombination is not a requirement for exon shuffling.28 Mutations in already existing genes without gene duplication In this process gradual accumulation of various mutations changes the structure and function of the original gene. In addition, genes can also gain new functions, without gene duplication, by changes in their regulatory elements14,29 and by differential splicing.30

Gene Networks and Supernetworks

375

Evolution of Paralogous Genes Redundant genes are prone to being lost from the genome, as there may be no selection pressures to retain them. So what is the reason that, after gene or genome duplication, the extra copies of any specific gene are retained in the genome and not eliminated or turned into pseudogenes? At least two answers can be given. One is that the extra copies of the same gene have been retained because it is advantageous to have a backup system. If there would be a deleterious mutation in one gene copy, the sister copies can replace it. Examples of gene knockouts which do not result in any obvious phenotypic change have accumulated during the past few years (for example, see ref. 31). The simplest explanation for a lack of obvious phenotypic change is that a homolog of the gene that was knocked out has an overlapping expression domain, or that it is induced in the same domain after the knockout occurs, and can take over the function of the knocked out gene. It should be noted that the protective effect of a redundant gene may not generate sufficient selection pressure to retain it in the genome. However, a minute positive pressure may be all that is necessary.32 Genetic models indicate that gene redundancy is evolutionary stable if one of the redundant genes performs the same function with a lower efficacy.33 The second answer is that the duplicates of a gene are retained because they diverge structurally and acquire a different function and participate in a different, albeit similar, gene network. Many studies have shown that paralogous genes can have different developmental roles (for example, see refs. 22, 34-37). Once a paralog has gained a new function, it will become a more stable member of the genome. It is likely that both of the above answers are correct. Paralogous genes often have overlapping expression domains and can substitute for each other if a mutation occurs in one. At the same time they may have different functions and participate in different gene networks. By comparison of the expression pattern of paralogous genes one can gain insight into the expression and function of the ancestral gene. The overlapping area of the expression of two paralogs is probably the site where the ancestral gene was expressed.38 After duplication, the paralogous genes maintain their expression and function in their ancestral site. However, later they may acquire new enhancer elements which direct their expression in additional locations and/or times. In the new environment(s), they may gain functions which are different than their ancestral one. After a long time, each gene may acquire specific functions that are different from those of its paralogs (for example, see refs. 39, 40).

A Gene Usually Participates in Many Different Networks If we look at the expression of various genes during development of members of a species, we find that the same genes are expressed at different times and locations. One way to interpret such expression patterns is that each gene is active in the same network, which functions in different locations and times in a continuous manner, and that throughout the time that the gene is being expressed, it has but a single and constant function. Another interpretation is that such genes function in different networks and have different roles. The analysis of expression pattern of many genes involved in development indicates that most are expressed many times during development in different locations.16 The same conclusion could also be drawn from enhancer trap experiments which show that most genes are expressed at different sites and different times.16 Although expression of a gene at a specific site does not automatically mean that the gene has a functional role in that location,41,42 many knockout studies show pleiotropic phenotypic changes in which the affected individuals display defects in multiple sites.43,44 The simplest way of interpreting multiple defects as a result of a gene knockout is that the gene has a functional role in the affected areas. However, in order to see if the gene participates in different developmental networks at

376

Molecular Basis of Epithelial Appendage Morphogenesis

different locations, one has to look closely at the function of the gene at each location. Such observations usually reveal that genes function in more than one network (for example, see refs. 35, 45, 46). To gain more insight into function of a gene, interspecies comparison will be highly useful.47,48 As will be discussed later, paralogous genes found in individuals of the same species may participate in different gene networks. Such gene networks are homologous and have gene members which are paralogous. However, in some cases they share the same gene. Although in each gene network the gene may have the same molecular function, the function of each network may be slightly different. There is a possibility that during evolution such genes may undergo a duplication event.49 The duplicated copies can then participate in different gene networks, later fine-tuning their function for their specific role in their specific network (for example, see ref. 50).

Orthologous Genes Usually Have Similar Functions During the past decade it has been found that the development of many homologous structures in various species is regulated by genes that are themselves orthologous. The following is in support of the premise that orthologous genes found in different species function very similarly to each other: 1. Often the expression of orthologous genes is very similar among closely and distantly related species (for example, see refs. 38, 51-55). 2. Knockouts of the orthologous genes usually lead to similar phenotypic defects (for example see refs. 56, 57). 3. In certain cases, orthologous genes can substitute for each other with no apparent change in phenotype (for example, see refs. 58, 59). 4. Regulatory regions of orthologous genes are usually conserved and can substitute for each other with only minor changes in expression patterns.60 The conservation of expression and regulation of orthologous genes indicate that their role is most likely conserved among various species. As will be mentioned below, since proteins function by directly or indirectly interacting with other proteins or genes, it is very likely that such interactions have also been conserved.

New Gene Expression Patterns Are Acquired Through Changes in the Enhancer Sequences During evolution, the expression of a gene can be modified by changes in their enhancer sequences.61 Enhancer sequences can change by simple changes in the enhancer elements, such as point mutations and small deletions and additions. Larger deletions or additions can have more drastic effects, as they may affect more than one enhancer element. Gene shuffling where an entire gene is transferred from one location in the genome to another may entirely change the expression pattern of a gene. Enhancer sequences can also be shuffled around and may land upstream or downstream of a gene, changing its expression pattern. Generally changes in enhancer sequences can have either qualitative or quantitative consequences on the expression of a gene: Qualitative changes Heterochronic changes These changes modify the timing of the expression of a gene. Thus a gene that is expressed during one period of development will be expressed in another period. It should be noted that heterochronic change as described here is different from the definition given to it

Gene Networks and Supernetworks

377

previously, where it is described as relative shifts in timing of expression of certain anatomical features in a certain species.62-64 Heterotopic changes These changes modify the site of the expression of a gene. Thus a gene that is expressed in one specific location will be expressed in another location. Quantitative changes These changes modify the degree of the gene expression. Thus a gene that is expressed at a certain level, will be expressed at a higher or lower level.12,65 Each or a combination of the above can serve as a mechanism whereby the expression pattern of a duplicated gene or gene network can be changed. For example, changes in the expression pattern of one or a few genes within a specific network can change the expression of the whole network (see Fig. 19.1).

Conservation of Gene Interaction Whether they are structural or functional, proteins interact with other proteins either directly or indirectly. Direct interactions are those in which two or more proteins physically interact with each other by directly binding to each other and changing each other’s activity or availability. Indirect interactions are those in which one protein modulates the transcription, translation, post-translational modification, availability, and various nonprotein interaction(s) of another protein. Conservation of gene structure and function is indicative of conservation of protein interactions. This is another reason that de novo appearances of new genes are rare. During their evolution, genes usually acquire a function within the context of the other genes with which they interact. Once a network of genes that interact with each other and have specific functions is established, a restriction would be placed on any one gene changing its function within the network.12,66 A change in the function of a gene is only allowed by coordinated compensations in the function of other members of the network, and therefore occurs with a low probability. Thus a network of interacting genes is an evolutionarily stable system as long as it has an adaptive advantage. Of course, many structural changes in genes are still tolerated as long as they do not affect the interactional integrity and function of the network.47 The property of a gene network restricting changes in the function of its members is an example of an evolutionary constraint.67,68 A well known example of an evolutionary constraint due to the strict conservation of gene interactions may be the phylotypic stage.69 The phylotypic stage is defined as the stage of development at which members of a phylum look most similar to each other and the position of all the major body parts has been determined.66,69,70 For vertebrates, the phylotypic stage is the tailbud stage, for nematodes it is the stage after the completion of most embryonic cell divisions and for arthropods it is the fully segmented germband stage. Although conserved gene interactions occur both before and after this stage, at this stage the gene networks that are active are most similar to each other, as indicated by the similarity of body plans. Phylotypic stage could be compared to the foundation of, for example, a building. One could make many different buildings that look different from outside, but use the same standard structural foundation. Although the phylotypic stage itself is not entirely conserved,71 the conservation and constraint at this stage may serve as a solid foundation on which evolution works to bring about diversification of body plans.72 Interestingly, the phylotypic stage is correlated with the sequential expression of Hox genes, which may indicate an important role for this gene family in establishment of this stage.73

378

Molecular Basis of Epithelial Appendage Morphogenesis

Conserved gene interactions do not always give rise to homologous morphologies. Sometimes a specific gene network may be involved in two processes which give rise to completely different structures. The homology between the two structures is called the homology of process.74 Gene networks which cause homologies of process can be found by comparing different structures in the individuals of the same species. For example, it appears that in vertebrates, close similarities exist in the processes that occur in the morphogenesis of various structures that arise through epithelial-mesenchymal interactions.75-77 Although the gene interactions in these secondary inductive events are similar, the structures that arise from them are quite varied. Thus in the same individual, the process of morphogenesis of a tooth may be similar to that of a fingernail, although the resulting structures are very different. Gene networks that cause homologies of process can also be found by comparing individuals of different species. This can be illustrated by the concept of zootype, which is defined as the specific “spatial pattern of gene expression” that is characteristic of each species.66 Thus, for example, in closely related species, the genes and gene expression are very similar, except that slight differences have appeared among these species regarding their specific spatial pattern of gene expression. Here we would like to emphasize that the timing and quantity of gene expression is also important. Thus each species has its own unique qualitative (i.e., spatiotemporal) and quantitative way of expressing genes. During the phylotypic stage, these differences between members of a phylum are minimal. However, it is very likely that many of the gene interactions involved in various aspects of development, before and after the phylotypic stage, are still similar but the spatiotemporal patterns of gene expression are unique, thus in each species giving rise to individuals with quite distinct development and morphologies. In spite of the above, it is quite possible that differences in development of various species are also a result of changes in the connectivity of genes, i.e., changes in the circuitry of gene networks78 or changes in the function of regulator genes.79 Although such changes would occur with lower likelihood, they may contribute more significantly to differences in the developmental programs (see the next section). In the next section we will present the gene network and supernetwork models. Our purpose in presenting these models is to consolidate some basic observations about developmental programs and the genes that are responsible for them. Many of the developmental programs found among and within various species are derived from older programs that were duplicated and modified. The fact that these programs are similar can be deduced from the observation that the genes and gene interactions responsible for them are similar.

Gene Networks Definition of a Gene Network A gene network can be defined as an association of many genes which interact with each other in cascades or parallel pathways and achieve a specific function or various functions through such interactions. The members of a gene network usually can directly or indirectly modulate each other’s expression or function through stimulation or inhibition of transcription, translation or post-translational modifications. Each gene network usually regulates a specific function during development. For example, a gene network may regulate the anterior-posterior polarity and another the length of an organ. Some gene networks may regulate more than one function. The gene members within a multifunctional gene network may have complex regulatory elements directing their expression to various spatiotemporal environments.

Gene Networks and Supernetworks

379

Conservations of Gene Networks Many genes in nature are not only evolutionarily conserved in their structure but also in their interaction with other genes. The way genes in a network interact with each other we call gene interaction circuitry. Gene networks are under tight evolutionary constraint not to change their circuitry and function. Similarly to genes functioning within the context of other genes, gene networks also function in the context of other networks (see gene supernetworks below). Because of this, any modifications that disrupt the function of a gene network may lower the fitness of an individual or be deleterious. Thus gene networks are usually stable systems and would be conserved during evolution and speciation. Variations to a gene network can be tolerated as long as they do not disrupt the preexisting network. For example, branching in or duplication of the network are well tolerated. Branching is defined as new pathways that originate from somewhere within the preexisting network and do not disturb the already established function of the network. Branching can lead to a gain of new function, which may or may not be manifested in the formation of a new structure. Duplication is defined as the formation of a new copy of the entire gene network, when the new network can function similarly to the original network.

Duplication and Divergence of Gene Networks Gene networks can be duplicated by the following two processes: Polyploidization It is very likely that most of the similar gene networks that exist today arose through genome duplication. Once a genome duplication occurs, the resulting daughter gene network can function identically to the parental network by reestablishing its original interaction and resuming its original function. Subsequently over time, the two copies of the gene network may slightly change their function by modifying some of their genes’ structure and interactions. The process of the duplication of a gene network is thus a simple one if genomes are duplicated. Individual gene duplications There is also the possibility that a gene network is duplicated by individual gene duplications through duplication of short stretches of DNA. In such a case a different duplication event is necessary for each gene in order to duplicate the entire network if the genes are located on different chromosomes. However, if the genes in the network are located close to each other on one chromosome, only one or two duplication events involving long stretches of DNA may suffice. It appears that the most favorable situation is when the genes in a network are either most closely linked or as far apart as possible.80 For gene networks that contain genes that are located on different chromosome, the process can be broken down into many steps of individual gene duplication (Fig. 19.2). Let us imagine that a specific gene network contains 5 distinct genes (in reality this number may be closer to 50 to 200). As soon as one of those genes is duplicated, a new network is established that can function similarly to the original network. Of course, 4 genes in each network are identical. Now over time, some of those 4 genes are also duplicated and can substitute for their counterparts. Eventually all or a fraction of the genes may be duplicated and thus a duplicated gene network is established that can function exactly as its homolog. After each gene duplication, the duplicate copies could each be shared by both networks. Additionally, there are some genes that may never undergo a duplication event and therefore would participate in both networks. If a deleterious mutation occurs in one of the gene members, the function of the old network is not disrupted, since the paralog of the mutated gene can substitute for it. On the other hand, instead of being deleterious, the mutation can

380

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 19.2. Duplication of a hypothetical gene network by genome or individual gene duplication. Each geometric figure represents a specific gene. The unfilled figures are duplicated versions of and exactly the same as the original genes (shown in black). Genome duplication results in two gene networks that have the exact genes, interactions, and function. One of the duplicates may later diverge structurally (by gain of function mutation in its gene members or changes in the circuitry of the network) and gain a new function. Over long evolutionary periods, the function of one of the duplicated networks may become completely different from its ancestral network. However, the two networks may still retain enough similarity to be recognized as homologous. Duplication of a gene network can also occur by duplication of short stretches of DNA containing a specific gene. In steps 1, 2, and 4 only one gene is duplicated, while in step 3 two genes are duplicated due to close linkage. It should be noted that complete duplication of all the members of a gene network is not necessary to establish a new network. This is due to the participation of one or more unduplicated genes in both networks.

be a gain of function mutation which causes, for example, a branching or a new connection in the network. It should be noted that there is a finite time (0.5-6 million years) during which a duplicated gene should acquire a function and become a permanent member of the genome before acquiring many mutations that may make it a nonfunctional gene.81 One can imagine that mutations can occur in one of the duplicate copies so that a new gene network is formed that now has a different function from the original one. Although different members of both gene networks are still homologous in their gene sequence and interactions and have roughly similar functions, each gene network works a little differently than the other. Over long periods of time, a duplicated gene network may gain new functions, and evolutionarily diverge in such a way that it does not share very close similarities with its ancestral function.

Gene Networks and Supernetworks

381

Paralogous and Orthologous Gene Networks Homologous gene networks can be distinguished from each other based on the identity of their genes and the interactions among the gene members (see Fig. 19.1). The gene members of two homologous gene networks have similar gene structure and the way their genes interact with each other (or their circuit design) is similar. Gene networks can be classified as either paralogous or orthologous. Paralogous gene networks are found in the individuals of the same species and created as a result of polyploidization or individual gene duplication, as described above. If one compares various gene members of the two paralogous gene networks, one would find some of the members are each other’s paralogs. Additionally, as mentioned before, the same gene may participate in two or more paralogous gene networks, having similar function(s) in each network. If this gene undergoes one or more duplication events, the duplicate copies may diverge over time to better adapt to their specific function in each paralogous network.49 Paralogous networks may be found to work in the same, overlapping, or different spatiotemporal domains in the individuals of the same species.82,83 Orthologous gene networks are found in individuals of different species84 and produced as a result of speciation. Many examples of paralogous77,85-88 and orthologous35,89-95 gene networks can be found in the literature. It should be noted that the task of identifying two similar gene networks as orthologous may sometimes be error prone. Although two gene networks in two different species may be homologous (i.e., derived from the same ancestral network), they may not be orthologous (i.e., having the closest ancestry). This is shown in Figure 19.3. Duplication of a gene network (GNo) in an ancestral species (So) results in two gene networks, GN1 and GN2, which have become evolutionarily diverged in both structure and function, but retain some similarity. Two descendants of this species, S1 and S2, both have GN1 and GN2. However, in S1, GN2 may be eliminated while in S2, GN1 may be eliminated. If now one compares GN1 in S1 and GN2 in S2, one may get the erroneous conclusion that GN1 and GN2 are orthologs. Thus when two gene networks are found in two different species that seem to be similar, it can not be automatically assumed that they are orthologous. In order to become more certain that two networks are orthologous, one has to look for the presence of other similar gene networks in the two studied species. Ultimately, the study of the gene network family in a few different species would make the matter more clear. Additionally, two gene networks may be orthologous but have slightly different gene interaction circuitry. For example, a species S1 may have a gene network, GNOrtho1, that is orthologous to GNOrtho2 in species S2. In both of the gene networks, one of the gene members, gene a (ga), stimulates the expression of a second gene, gb. However, during the evolution of S1, gb loses its ability to regulate the expression of ga but acquires the capability to direct the expression of another gene, gc, while such a change does not occur for S2. Thus although GNOrtho1and GNOrtho2 are orthologous, their gene interaction circuitry is slightly different (Fig. 19.4). Again, interspecies comparisons would have to be carried out to determine the orthology of the two gene networks.

Gene Supernetworks Definition of a Gene Supernetwork We have described the evolution and conservation of genes and gene networks. Now we would like to propose a higher level of assembly of genes, a gene supernetwork. A gene supernetwork is defined as a collection of gene networks which participate with each other during the morphogenesis of a specific structure, for example an organ, a segment, or an appendage. Gene networks within a supernetwork can interact with each other by modifying the transcription, translation, availability or activity of the gene members of the other

382

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 19.3. Two nonorthologous gene networks may be erroneously identified as orthologous. A hypothetical species (So) has a gene network (GNo) which undergoes a duplication event, giving rise to GN1 and GN2. Over time these gene networks diverge both structurally and functionally but still retain some similarity. Later this species undergoes speciation giving rise to species 1 (S1) and species 2 (S2), both retaining the two gene networks. During evolution, GN2 is lost from S 1 while GN1 is lost from S2. Now if one compares GN1 and GN2 with each other, one may erroneously conclude that these networks are orthologous. Thus establishment of orthology requires closer inspection of the gene members, gene interactions, and the function of networks. This may be accomplished by comparing these networks in a few different species.

networks (Fig. 19.5). Each network within a supernetwork has its own unique set of genes, circuitry and specialized function, and thus can be easily distinguished from other networks. However, it is also possible that two or more paralogous networks are functional within the same supernetwork. It is also possible that the same gene may participate in more than one network within a supernetwork, having similar or slightly different roles in each network. Additionally, the same gene network may function within two or more different supernetworks. Indeed it is quite possible that supernetworks active within any species are assembled from a limited number of gene networks. Thus a few dozen gene networks combine in various combinations to give rise to a few different gene supernetworks.

Duplication, Divergence and conservation of a Gene Supernetwork As with gene networks, gene supernetworks can be duplicated during evolution, thus giving rise to new structures which are the same as or different from the original structure. When a gene supernetwork is duplicated, the duplicate is not under the same evolutionary pressure as its old counterpart and thus can be modified so that it can give rise to a new structure. For example, a gene supernetwork for morphogenesis of a limb can be duplicated, and later one of the duplicates may undergo heterotopy so that it can give rise to a new limb in a different location. Later in evolution, this supernetwork can go through further changes in its constituent gene networks so that it may give rise to a structure which is phenotypically quite different from a limb. Many structures present today which may not be morphologically similar may have the same evolutionary origins. By studying the molecules involved in the morphogenesis of such structures, one can unmistakably determine if they are homologous or not.96

Gene Networks and Supernetworks

383 Fig. 19.4. A difference in the circuitry of two gene networks may not indicate a lack of orthology between the networks. Two hypothetical species, S1 and S2 which have the same ancestry, each have a gene network which is orthologous to the other. The two gene networks, GNortho1 and GNortho2, have many orthologous gene members but the focus will be on only the two genes which are shown, gene a and gene b. The arrows indicate that each gene stimulates the expression of the other gene. Thus both gene a and gene b can stimulate the expression of the other. During the evolution of Gnortho1 in S1, gene b loses its ability to stimulate the expression of gene a, but gains the ability to stimulate the expression of another gene, gene c. No such change occurs for GNortho2. Since Gnortho1 and Gnortho2 have a different circuitry and interaction, one may erroneously conclude that the two networks are not orthologous. Again, multi-species comparisons may clear the matter.

Due to their crucial role in morphogenesis, gene supernetworks, like gene networks, are conserved during evolution and speciation. This has been especially supported by molecular studies during the development of various animal limbs.97 For example, gene supernetworks that are responsible for the morphogenesis of fly and vertebrate limbs,55,98 fore and hind limbs of land-dwelling vertebrates,85-87 and fly limb and wing88 are very similar, indicating a very strict evolutionary conservation of these supernetworks. Experiments with epithelial-mesenchymal recombination among two distantly related species, mouse and chicken,99 and closely related species, newt and frog,100 indicate that mechanisms involved in the morphogenesis of appendages are partially conserved among these species. This argues for the possibility that the gene supernetworks involved may be also partially conserved. Even in appendages as different as teeth and feathers, supernetworks giving rise to them appear to be at least partially conserved. Many molecules expressed during the induction of feather and teeth are very similar. For example, at a certain stage of early development, the expression patterns of Msx genes in teeth101 and feathers102 correspond to each other. Indeed many molecules which are expressed during the induction of feather, hair, limb, and tooth appear to be identical, and thus point to the possibility that these appendages are a result of the same ancestral programming.

384

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 19.5. A hypothetical gene supernetwork. Each numbered black circle represents a different gene. Each circle represents a gene network. This supernetwork is made up of five different gene networks. A real and typical supernetwork may be composed of many more gene networks. Gene networks contained in a supernetwork can modulate each other’s activity. The sharp and blunt arrows indicate that the upstream gene is respectively stimulating or inhibiting the activity, availability, expression, or translation of the downstream gene through direct or indirect protein interactions. Not shown here is the possibility that a supernetwork may contain two or more gene networks that are paralogous to each other. In addition, different gene supernetworks may share some of the same gene networks. Note that if the boundaries (the large circles) around each gene network are taken away, the supernetwork is transformed into a giant network of genes. In reality there are no boundaries between the genes and gene networks within a supernetwork. However, a group of genes which cooperate and interact with each other to perform a specific function within a supernetwork, we call a gene network. Distinctions between gene networks have to be made by close examination of the function of each network and by interspecies comparisons.

Differences in Development Among Various Species Is Due to Differences in Their Gene Networks and Supernetworks, Not in Their Genes The differences between two closely related species will not be clear if we look at their genes, but become more clear as we look at their gene networks and supernetworks. From identification of many orthologous genes among species, it appears that most animals, even if they are not closely related, have the same types of genes. Closely related species have been found to possess very similar genes.19,103-105 So why is it that different species have such different morphologies? A simple answer would be that the differences reside in the regulatory regions of the genes. Thus although the coding regions of genes are similar and have similar function, their regulatory regions which determine their qualitative and quantitative expression patterns are different among species. Such differences cause genes to be expressed differently in different species, thus giving rise to differences in morphology. The differences are small among closely related species, but large among distantly related ones. Additionally, in direct correlation with changes in the regulatory regions may be changes in regulatory genes which control the expression of other genes.79 There are two ways that changes in regulatory regions can cause differences in morphology among species: Modifications which do not change gene network and supernetwork circuitry Such modifications only change either the quality (i.e., heterochronic or heterotopic changes) or quantity of gene expression without establishing new connections between various genes. Thus the circuitries of the networks and supernetworks are not changed. To give a hypothetical example, gene x in network A which induces the expression of gene y in network B still does so, except that the time, location, or quantity of this induction is different than before.

Gene Networks and Supernetworks

385

Modifications which change gene network and supernetwork circuitry Such modifications cause old connections to be disrupted or new connections to form among various genes which function in different gene networks and supernetworks. For example, gene x in gene network A that previously did not regulate the expression of gene y in network B, does so now. From most of the evidence gathered so far, it appears that many orthologous gene networks have conserved gene interaction circuitries. At the present time, the same is not determined for gene supernetworks and thus one could not tell which of the above alternatives is correct. It is very likely that both of the above mechanisms are involved in establishing differences in morphology among species. However, many changes in the morphology of two closely related species can be explained by qualitative and quantitative changes rather than changes in the circuitry of gene networks and supernetworks. For example, let us suppose that a gene supernetwork is involved in the development of a skin appendage in two different species. Two gene networks working within this supernetwork are respectively responsible for determination of the length and the width of the appendage. It would be easy to imagine that if a heterochronic change would occur so that in one species both of the networks are working for a longer time, both the length and the width of the appendage would increase in that species, leading to a change in morphology. Over time such small changes would lead to drastic modifications of a species’ morphology. In very closely related species (i.e., mouse and rat), the gene supernetworks may be so similar that the reason for a morphological change may not be obvious. Thus the total assembly of the gene supernetworks have to be considered. Here we present another term: gene totinetwork. A gene totinetwork is an assembly of many gene supernetworks and capable of generating a whole organism. Thus it is totipotent. The hierarchy of gene organization can be defined for each member of a species by the following: Genes → Gene Networks → Gene Supernetworks → Gene Totinetwork Thus each species, even if it is closely related to another, has its own unique gene totinetwork. Gene totinetwork is different from zootype. Zootype is defined as the specific “spatial pattern of gene expression” that is characteristic of each species.66 A gene totinetwork, on the other hand, is a functional unit which is composed of genes that have a hierarchical organization, being made up of many gene supernetworks. Each supernetwork is responsible for making a part of a whole organism. Unlike gene networks within a supernetwork which are interactive, gene supernetworks within a totinetwork are more independent of each other. They may only interact with each other during phylotypic stage69 and/or regeneration and repair. The independence of supernetworks can also be deduced from the concept of dissociability.106 Comparison of various species indicates that the timing of various events during development can be varied independently. For example, the time when limb buds, somites, heart tubes, nasal placodes, and heart placodes appear in various species is different from each other.71 This may indicate that during the evolution of various species, gene supernetworks involved in the generation of the above structures have undergone heterochronic changes in their expression. Thus each supernetwork acts as an independent module,69,107 which by itself is sufficient to generate a specific structure and can undergo qualitative and quantitative shifts in its expression.

Can an Amphibian Limb Be Converted to a Fish Fin with Genetic Manipulation? If one knows about the circuitry and expression pattern of gene networks within a supernetwork, it would be possible to manipulate, for example, a limb supernetwork in

386

Molecular Basis of Epithelial Appendage Morphogenesis

such a way that it would give rise to a fin (or vice versa). Along the same lines, gaining a complete knowledge of how a limb supernetwork works, which may not be an easy task,108,109 will give us the ability to make other limbs which look very different.

Can an Inactivated Gene Supernetwork be Reactivated? During the evolution of a species, a gene supernetwork that is responsible for the formation of, for example, a limb, finger, feather, tail, or tooth may become inactivated and thus the corresponding structure cease to form. However, other paralogous supernetworks in that species may contain many of the same gene networks once possessed by the inactivated supernetwork. Thus there is a possibility that the supernetwork can be reestablished by reassembly of all the gene networks which it once contained. Such reassembly may be achieved simply by heterochronic, heterotopic, or quantitative shifts in the expression of some key genes contained within the gene networks (for example, see refs. 110-112). On the other hand, an inactivated supernetwork that has a unique gene network, not shared by any other network, can become reactivated only during a short time (0.5-6 million years)81 before mutations disrupt the functioning of the unique gene network.

Summary and Conclusion Although genes evolve as individual units, they function in networks involving many genes. During the past two decades, it has become increasingly clear that in order to fully understand the function of a specific gene, one has to look at its interactions with other genes. Interestingly, it has been found that many genes involved in development not only have evolutionarily conserved structures but have conserved gene interactions. A group of genes that work together to achieve a specific function during development is called a gene network. A change in the function of a gene network can be achieved if some or all members of the gene network are duplicated (both promoter and coding sequences) and an extra copy of the gene network is established. The extra copy now can functionally diverge when its gene members have gain of function mutations which cause changes in gene interactions (circuitry) and expression (i.e., changes in time, site, and quantity of expression). The duplication and divergence is probably a slow process because each gene functions in the context of other genes, and thus structural and functional changes in one gene affect the whole network. The function of a new gene network may be similar or different than its ancestral function, depending on the length of time since its duplication and divergence. Gene networks which have the same ancestor can be distinguished from others because they have gene members with conserved sequence and position in the network. The duplication of gene networks is complicated by the possibility that individual members of the network may be located on different chromosomes. Many gene networks, having the same or different ancestry, can participate with one another to create a gene supernetwork. The function of a gene supernetwork is more complicated than a gene network. For example, a gene network may be involved in a process like cell proliferation during the development of a fish fin. But a gene supernetwork participates in all of the different aspects of the development of the fin. Gene supernetworks can in turn undergo duplication and divergence. For example, a gene supernetwork that functions in construction of a amphibian scale can give rise to a supernetwork that can construct an avian feather. As with gene networks, the basic unit of change in a gene supernetwork is a gene. Thus a change in one gene may affect the entire supernetwork. During evolution new structures may have formed by new associations between gene networks and a supernetworks. For example, it could be imagined that a gene supernetwork for constructing reptile scales was linked to a network that directs proximal distal growth. This association could have transformed scales into elongated structures, the precursors of

Gene Networks and Supernetworks

387

hairs or feathers. It is also possible that such a supernetwork may branch to incorporate a gene network responsible for compartmentalization to form alternating epithelial stripes which subsequently undergo keratinization and cell death (see chapter 5). It is further possible that by simply activating a similar homologous gene network, in a fractal like fashion, barbules can form on top of barbs. The product is a three-level-branched skin appendage, fluffy and air trapping, ideal for keeping organisms warm—a big evolutionary advantage for the dinosaur/the bird. Finally, by invoking another gene network that makes barbule cells form hook-like structures, the numerous barbules and barbs are knit into a strong plane-like appendage that allow the birds to fly—an even bigger evolutionary advantage that opens the sky to the bird. The result of this successful assembly of gene networks is an “evolution patent” that rewards this gene supernetwork with 88,000 products (or bird species). As can be deduced from the conservation of developmental genes and their interactions, the mechanisms of evolution are mostly conservative.113 New genes come from old genes, new interactions come from old interactions and new functions come from old functions. In this chapter we did not address the possibility that similarities in developmental programs may arise through convergent evolution. This possibility may be true in a minority of cases due to the constraints in developmental systems which may predispose different species to acquire the same genetic programming.69 However, it should be noted that since the possession of such constraints is due to shared ancestry, acquisition of the same genetic program is really a case of parallel evolution, not convergent evolution.65 Understanding of a gene supernetwork, for example, involved in the development of a limb or a skin appendage, will enhance our task of understanding other supernetworks. However, it is not an easy task to identify all of the genes that function within a gene supernetwork. Additionally, it is an even more challenging job to identify all of the gene interactions. Even if we know all of the genes and their interactions, we still may not be able to easily understand the properties and the function of a supernetwork.109 This has to do with the complexity of the system and the possibility that if we study the isolated components of a system, we may not be able to understand the property of the whole system. However, once a few key principles are understood, a thorough understanding may not be necessary in order to modulate the function of a gene supernetwork. Such modulation would benefit us in regrowth of hair, an amputated limb, or an inactive or faulty organ, and later may enable us to engineer novel limbs or organs.

Acknowledgment This work was supported by grants from the NIH and the NSF to C-M. C. and V.H. We thank Joe Staton, Judith Lengyel, David Jacobs, Ann Daniel, and Karin Dumstrei for helpful comments on the manuscript.

References 1. Doolittle RF. Reconstructing history with amino acid sequences. Prot Sci 1992; 1:191-200. 2. Green P. Ancient conserved regions in gene sequences. Curr Opin Struc Biol 1994; 4:404-412. 3. Dayhoff MO. Computer analysis of protein sequences. Fed Proc 1974; 33:2314-2316. 4. Doolittle RF. Similar amino acid sequences: chance or common ancestry? Science 1981; 214:149-159. 5. Dorit RL, Schoenbach L, Gilbert W. How big is the universe of exons? Science 1990; 250:1377-1382. 6. Chothia C. One thousand families for the molecular biologist. Nature 1992; 357:543-544. 7. Chothia C. Protein families in the metazoan genome. Development Supplement 1994:27-33.

388

Molecular Basis of Epithelial Appendage Morphogenesis

8. Green P, Lipman D, Hillier L et al. Ancient conserved regions in new gene sequences and the protein databases. Science 1993; 239:1711-1716. 9. Blundell TL, Johnson MS. Catching a common fold. Prot Sci 1993; 2:877-883. 10. Orengo C. Classification of protein folds. Curr Opin Struct Biol 1994; 4:429-440. 11. Zuckerkandl E. The appearance of new structures and functions in proteins during evolution. J Mol Evol 1975; 7:1-57. 12. Zuckerkandl E. Topological and quantitative relationships in evolving genomes. In: C Helene, ed. Structure Dymanics Interactions and Evolution of Biological Macromolecules. Dordrecht, Holland: D Reidel Publishing 1983:395-412. 13. Murzin A. Can homologous proteins evolve different enzymatic activities? Trends Biochem Sci 1993; 18:403-405. 14. Piatigorsky J, Wistow G. The recruitment of crystallins: New functions precede gene duplication. Science 1991; 252:1078-1079. 15. Ohno S. Evolution of Gene Duplication. Heidelberg: Springer-Verlag, 1970. 16. Miklos GLG, Rubin GM. The role of the genome project in determining gene function: Insights from model organisms. Cell 1996; 86:521-529. 17. Sidow A. Gen(om)e duplication in the evolution of early vertebrates. Curr Opin Genet Dev 1996; 6:715-722. 18. Holland PW, Garcia-Fernandez J, Williams NA, Sidow A. Gene duplications and the origins of vertebrates. Development Supplement, 1994; 125-133. 19. Lundin LG. Evolution of the vertebrate genome as reflected in paralogous chromosomal regions in man and the house mouse. Genomics 1993; 16:1-19. 20. Schughart K, Kappen C, Ruddle FH. Duplication of large genomic regions during the evolution of vertebrate homeobox genes. Proc Natl Acad Sci USA1989; 86:7067-7071. 21. Moon RT. In pursuit of the functions of the Wnt family of developmental regulators: Insights from Xenopus laevis. Bioessays 1993; 15:91-97. 22. Hogan BLM. Bone morphogenetic proteins: Multifunctional regulators of vertebrate development. Genes Dev 1996; 10:1580-1594. 23. Coulier F, Pontarotti P, Roubin R et al. Of worms and men: An evolutionary perspective on the fibroblast growth factor (FGF) and FGF receptor families. J Mol Evol 1997; 44:43-56. 24. Patthy L. Modular exchage principles in proteins. Curr Opin struct Biol 1991; 1:351-361. 25. Doolittle RF, Bork P. Evolutionary mobile elements in proteins. Sci Am 1993; 269:50-56. 26. Doolittle RF. The multiplicity of domains in proteins. Annu Rev Biochem 1995; 64:287-314. 27. Patthy L. Introns and exons. Curr Opin Struct Biol 1994; 4:383-392. 28. Patthy L. Exon shuffling and other ways of module exchange. Martix Biol 1996; 15:301-310. 29. Palopoli MF, Patel NH. Neo-Darwinian developmental evolution: Can we bridge the gap between pattern and process? Curr Opin Genet Dev 1996; 6:502-508. 30. Lopez AJ. Developmental role of transcription factor isoforms generated by alternative splicing. Dev Biol 1995; 172:396-411. 31. Saga Y, Yagi T, Ikawa Y et al. Mice develop normally without tenascin. Genes Dev 1992; 6:1821-1831. 32. Tautz D. Redundancies, development and the flow of information. BioEssays 1992; 14:263-266. 33. Nowak MA, Boerlijst MC, Cooke J, Maynard Smith J. Evolution of genetic redundancy. Nature 1997; 388:167-171. 34. Kingsley DM. The TGF-β superfamily: New members, new receptors, and new genetic tests of function in different organisms. Genes Dev. 1994; 8:133-146. 35. Moon RT, Brown JD, Torres M. WNTs modulate cell fate and behavior during vertebrate development. Trends Genet 1997; 13:157-162. 36. Condie BG, Capecchi MR. Mice with targeted disruptions in the paralogous genes hoxa-3 and hoxd-3 reveal synergistic interactions [published erratum appears in Nature 1994 Oct 6;371(6497):537]. Nature 1994; 370:304-307. 37. Chen F, Capecchi MR. Targeted mutations in hoxa-9 and hoxb-9 reveal synergistic interactions. Dev Biol 1997; 181:186-196. 38. Holland P. Homeobox genes in vertebrate evolution. BioEssays 1992; 14:267-273.

Gene Networks and Supernetworks

389

39. Fromental-Ramain C, Warot X, Lakkaraju S et al. Specific and redundant functions of the paralogous Hoxa-9 and Hoxd-9 genes in forelimb and axial skeleton patterning. Development 1996; 122:461-472. 40. Sanford LP, Ormsby I, Gittenberger-de Groot AC et al. TGF-β2 knockout mice have multiple developmental defects that are non-overlapping with other TGF knockout phenotypes. Development 1997; 124:2659-2670. 41. Dickinson WJ. On the architecture of regulatory systems: Evolutionary insights and implications. BioEssays 1988; 8:204-208. 42. Erickson HP. Gene knockouts of c-src, transforming growth factor β1, and tenascin suggest superfluous, nonfunctional expression of proteins. J Cell Biol 1993; 120:1079-1081. 43. Parody TR, Muskavitch MA. The pleiotropic function of Delta during postembryonic development of Drosophila melanogaster. Genetics 1993 135:527-539. 44. Elson A, Wang Y, Daugherty CJ et al. Pleiotropic defects in ataxia-telangiectasia proteindeficient mice. Proc Natl Acad Sci USA 1996 93:13084-13089. 45. Artavanis-Tsakonas S, Matsuno K, Fortinin ME. Notch signalling. Science 1995; 268:225-232. 46. Schweitzer R, Shilo B-Z. A thousand and one roles for the Drosophila EGF receptor. Trends Genet 1997; 13:191-196. 47. Dover GA. Observing development through evolutionary eyes: A practical approach. BioEssays 1992; 14:281-287. 48. Patel NH. Developmental evolution: Insights from studies of insect segmentation. Science 1994; 266:581-590. 49. Hughes AL. The evolution of functionally novel proteins after gene duplication. Proc R Soc London Series B Biol Sci 1994; 256:119-124. 50. Kelly GM, Greenstein P, Erezyilmaz DF, Moon RT. Zebrafish wnt8 and wnt8b share a common activity but are involved in distinct developmental pathways. Development 1995; 121:1787-99. 51. Patel NH, Martin-Blanco E, Coleman KG et al. Expression of engrailed proteins in arthropods, annelids, and chordates. Cell 1989; 58:955-968. 52. Fietz MJ, Concordet JP, Barbosa R et al. The hedgehog gene family in Drosophila and vertebrate development. Development Supplement 1994:43-51. 53. Nagy LM, Carroll S. Conservation of wingless patterning functions in the short-germ embryos of Tribolium castaneum. Nature 1994; 367:460-463. 54. Davidson D. The function and evolution of Msx genes: Pointers and paradoxes. Trends Genet 1995; 11:405-411. 55. Panganiban G, Irvine SM, Lowe C et al. The origin and evolution of animal appendages. Proc Natl Acad Sci USA 1997; 94:5162-5166. 56. Noll M. Evolution and role of Pax genes. Curr Opin Genet Dev 1993; 3:595-605. 57. Hanson I, Van Heyningen V. Pax 6: More than meets the eye. Trends Genet 1995; 11:268-272. 58. Padgett RW, Wozney JM, Gelbart WM. Human BMP sequences can confer normal dorsalventral patterning in the Drosophila embryo. Proc Natl Acad Sci USA 1993; 90:2905-2909. 59. Zhao JJ, Lazzarini RA, Pick L. The mouse Hox-1.3 gene is functionally equivalent to the Drosophila Sex combs reduced gene. Gene Dev 1993; 7:343-354. 60. Cavener DR. Transgenic animal studies on the evolution of genetic regulatory circuitries. BioEssays 1992; 14:237-244. 61. Li X, Noll M. Evolution of distinct developmental functions of three Drosophila genes by acquisition of different regulatory regions. Nature 1994; 367:83-87. 62. Gould SJ. Ontogeney and Phylogeny. Harvard: Belknap Press, 1977. 63. Gould SJ. Ontogeny and phylogeny—Revisited and reunited. BioEssays 1992; 14:275-279. 64. Raff RA, Wray GA. Heterochrony: Developmental mechanisms and results. J Evol Biol 1989; 2:409-434. 65. Zuckerkandl E. Molecular pathways to parallel evolution: I Gene nexuses and their mophological correlates. J Mol Evol 1994; 39:661-678. 66. Slack JMW, Holland PWH, Graham CF. The zootype and the phylotypic stage. Nature 1993; 361:490-492.

390

Molecular Basis of Epithelial Appendage Morphogenesis

67. Alberch P. Developmental constraints in evolutionary process. In: Bonner JT, ed. Evolution and Development. Berlin: Springer-Verlag, 1982. 68. Maynard Smith J, Burian J, Kauffman S et al. Developmental constaints and evolution. Quart Rev Biol 1985; 60:265-287. 69. Raff RA. The Shape of Life: Genes, Development, and the Evolution of Animal Form. Chicago: The University of Chicago Press, 1996. 70. Wolpert L. The evolution of development. J Biol Linn Soc 1990; 39:109-124. 71. Richardson MK. Heterochrony and the phylotypic period. Dev Biol 1995; 172:412-421. 72. Gerhart J. Summing up: Conservation and diversification in metazoan eukaryotic cells. Phil Trans R Soc Lond B 1995; 349:333-336. 73. Duboule D. Temporal colinearity and the phylotypic progression: A basis for the stability of a vertebrate Bauplan and the evolution of morpholgies through heterochrony. Development Supplement 1994:135-142. 74. Gilbert SF, Opitz JM, Raff RA. Resynthesizing evolutionary and developmental biology. Dev Biol 1996; 173:357-372. 75. Burke AC. Epithelial-mesenchtmal interactions in the development of the chelonian Bauplan. Fortschr Zool 1989; 35:206-209. 76. Thesleff I, Vaahtokari A, Kettunen P, Aberg T. Epithelial-mesenchymal signaling during tooth develpment. Conn Tiss Res 1995a; 32:9-15. 77. Thesleff I, Vaahtokari A, Partanen A M. Regulation of organogenesis: Common molecular mechanisms regulating the development of teeth and other organs. Int J Dev Biol 1995b; 39:35-50. 78. Clarke B, Mittenthal JE, Senn M. A model for the evolution of networks of genes. J Theor Biol 1993; 165:269-289. 79. Hedrick PW, McDonald JF. Regulatory gene adaptation: An evolutionary model. Heredity 1980; 45:83-97. 80. Wagner A. Evolution of gene networks by gene duplications: A mathematical model and its implications on genome organization. Proc Natl Acad Sci USA 1994; 91:4387-4391. 81. Marshall CR, Raff EC, Raff RA. Dollo’s law and the death and resurrection of genes. Proc Natl Acad Sci USA 1994; 91:12283-12287. 82. Carroll SB. Developmental regulatory mechanisms in the evolution of insect diversity. Development Supplement. 1994:217-223. 83. Sander K. The evolution of insect patterning mechanisms: A survey of progress and problems in comparative molecular embryology. Development Supplement, 1994:187-191. 84. Nijhout HF. Symmetry systems and compartments in Lepidopteran wings: The evolution of a patterning mechanism. Development Supplement. 1994:225-233. 85. Gibson-Brown JJ, Agulnik SI, Chapman DL et al. Evidence of a role for T-box genes in the evolution of limb morphogenesis and the specification of forelimb/hindlimb identity. Mech Dev 1996; 56:93-101. 86. Simon HG, Kittappa R, Khan PA et al. A novel family of T-box genes in urodele amphibian limb development and regeneration: Candidate genes involved in vertebrate forelimb/ hindlimb patterning. Development 1997; 124:1355-1366. 87. Tabin C, Laufer E. Hox genes and serial homology. Nature 1993; 361:692-693. 88. Williams JA, Carroll SB. The origin, patterning and evolution of insect appendages. Bioessays 1993; 15:567-577. 89. Arendt D, Nubler-jung K. Common ground plans in early brain development in mice and flies. BioEssays 1996; 18:255-259. 90. de la Pompa JL, Wakeham A, Correia KM et al. Conservation of the notch signalling pathway in mammalian neurogenesis. Development 1997; 124:1139-1148. 91. De Robetis EM, Sasai Y. A common plan for dorsoventral patterning in bilateria. Nature 1996; 380:37-40. 92. Duboule D, Dolle P. The structural and functional organization of the murine Hox gene family resembles that of Drosophila homeotic genes. EMBO J 1989; 8:1497-1505. 93. Holland PWH, Garcia-Fernandez J. Hox genes and chordate evolution. Dev Biol 1996; 173:382-395.

Gene Networks and Supernetworks

391

94. Quiring R, Walldorf U, Kloter U, Gehring WJ. Homology of eyeless gene of Drosophila to the Small eye gene in mice and aniridia in humans. Science 1994; 265:785-789. 95. Salzberg A, Bellen HJ. Invertebrate versus vetebrate neurogenesis: Variations on the same theme. Dev Genet 1996; 18:1-10. 96. Jacobs DK. Developmental genes and the origin and evolution of metazoa. In: Schierwater B, Strait B, Wagner GP, DeSalle R, eds. Molecular Ecology and Evolution: Approaches and Applications. Switzerland: Birkhauser Verlag Basel, 1994. 97. Shubin N, Tabin C, Carroll S. Fossils, genes and the evolution of animal limbs. Nature 1997; 388:639-648. 98. Laufer E, Marigo V. Evolution in developmental biology: of morpholgy and molecules. Trends Genet 1994; 10:261-263. 99. Coulombre JL, Coulombre AJ. Metaplastic induction of scales and feathers in the corneal anterior epithelium of the chick embryo. Dev Biol 1971; 25:464-478. 100. Hamburgh M. Theories of Differentiation. New York: Elsevier, 1970. 101. Sharpe PT. Homeobox genes and orofacial development. Conn Tiss Res 1995; 32:17-25. 102. Noveen A, Jiang TX, Ting-Berreth SA, Chuong CM. Homeobox genes Msx-1 and Msx-2 are associated with induction and growth of skin appendages. J Invest Dermatol 1995 104:711-719. 103. King M-C, Wilson AC. Evolution at two levels in humans and chimpanzees. Science 1975; 188:107-116. 104. Wolfe KH, Sharp PM. Mammalian gene evolution: Nucleotide sequence divergence between mouse and rat. J Mol Evol 1993; 37:441-456. 105. Makalowski W, Zhang J, Boguski MS. Comparative analysis of 1196 orthologous mouse and human full-length mRNA and protein sequences. Genome Res 1996; 6:846-57. 106. Needham J. Biochemistry and Morphogenesis. Cambridge: Cambridge University Press, 1942. 107. Raff RA. Direct-developing sea urchins and the evolutionary reorganization of early development. Biol Essays 1992; 14:211-218. 108. Wolpert L. Do we understand development? Science 1994; 266:571-572. 109. Berkowitz A. Network of neurons, network of genes. Neuron 1996; 17:199-202. 110. Kurten B. Return of a lost structre in the evolution of the felid dentition. Societas Scientiarum Fennica, Commentationes Biol 1963; 26:3-12. 111. Kollar EJ, Fisher C. Tooth induction in chick epithelium: Expression of quiescent genes for enamel synthesis. Science 1980; 207:993-995. 112. Ledley FD. Evolution of the human tail. New Engl J Med 1982; 306:1212-1215. 113. Duboule D, Wilkins AS. The evolution of 'bricolage'. Trends Genet 1998; 14:54-59.

Part VI

Approaches

CHAPTER 20

Current Methods in the Study of Avian Skin Appendages Ting-Xin Jiang, N. Susan Stott, Randall B. Widelitz and Cheng-Ming Chuong

Introduction

C

hicken skin development is an excellent model to study the mechanisms of morphogenesis. It has a long experimental history and has been well characterized morphologically. Chicken skin offers distinct patterns and large numbers of different cutaneous appendages, accessibility to microsurgery, the availability of good in vivo and in vitro culture systems, and the presence of mutants to facilitate further investigation. In this chapter, we briefly describe current available methods, many used in our laboratory, to investigate the molecular pathways underlying the morphogenesis and maintenance of the chicken integument. Similar techniques are applicable to other epithelial appendages. A flow chart describing the experimental design used to analyze a new gene (Gene X) in our laboratory is shown (Fig. 20.1).

Detecting Molecular Expression To study molecules involved in feather morphogenesis, we first have to know what molecules are present within the tissue of interest at appropriate times. In situ hybridization and immunohistochemical methods detect mRNAs and proteins, respectively. These procedures are also useful to verify and characterize embryonic manipulations discussed below. For example, after viral perturbation it is essential to verify the transduction and to examine the altered expression patterns of other downstream genes. Therefore, these procedures are basic to molecular studies.

Whole Mount in Situ Hybridization with Nonradioactive Probes This method provides a three dimensional overview of the distribution of specific mRNAs, replacing tedious three dimensional reconstruction from stained sections using image analysis. In situ hybridization probes are easily generated from a small piece of skin (e.g., up to 50 mm3) by polymerase chain reaction (PCR).1 This technique has a large potential for mapping molecules involved in skin development in the future. Our laboratory uses a protocol based on that of Sasaki and Hogan.2 Briefly, skin from chicken embryos is dissected in RNase free phosphate-buffered saline (PBS) and fixed in 4% paraformaldehyde for 2 h or at 4°C overnight. Tissues are dehydrated, rehydrated, bleached, digested with proteinase K and then fixed again in 0.2% glutaraldehyde in 4% Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

396

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 20.1. Flow chart of the approaches to study the role of candidate molecules in epithelial appendage morphogenesis.

paraformaldehyde/PBS. Samples are then hybridized at 70°C overnight in hybridization buffer (Table 20.1) containing 2 µg/ml digoxigenin-labeled riboprobes. Samples are washed and treated with RNase A (50 µg/ml) before incubation with anti-digoxigenin Fab' conjugated to alkaline phosphatase (Boehringer Mannheim). Alkaline phosphatase is detected with 4.5 µl/ml 4-nitroblue tetrazolium chloride (NBT) with 3.5 µl/ml 5-bromo-4-chloro-3indolyl-phosphate (BCIP) (Promega) in color detection solution (Table 20.1) following standard protocols. The samples are then dehydrated, rehydrated, cleared and mounted. If desired, the sample can be embedded in wax and sectioned to identify the distribution of the probes and cell types with greater precision. For this purpose, the samples are dehydrated through methanol for 10 min, isopropanol for 15 min, and tetrahydronaphthalene for two 15 min washes and incubated three times in wax at 60°C for 20 min each wash. After the wax sets, sections are cut (7-20 µm), dried onto slides and then de-waxed with Histoclear.3

Current Methods in the Study of Avian Skin Appendages

397

Table 20.1. Formula for solutions 1. Hybridization Buffer 50% formamide 5 X SSC 1% SDS 50 µg/ml heparin 50 µg/ml tRNA 2. Color Detection Solution 100 mM Tris-HCl, pH 9.5 100 mM NaCl 50 mM MgCl2 0.5 mg/ml levamisol 0.1% Tween-20 3. Tris-glycine Buffer 24.2 g Tris base 15 g glycine DEPC-water to 2L Autoclave 4. Section In Situ Hybridization Buffer 40% formamide (Sigma Chemical) 5 x SSC 0.1 mg/ml sonicated herring sperm DNA (Life Technologies) 0.1 mg yeast tRNA 1X Denhardt’s solution (0.02% Ficoll, bovine serum albumin, polyvinyl pyrrolidone) 5. STE Buffer 500 mM NaCl 20 mM Tris-HCl, pH 7.5 1 mM EDTA

7. Blocking Solution 1% Blocking reagent (Boehringer Mannheim) 100 mM maleic acid 150 mM NaCl, Ph 7.5 8. Calcium-Magnesium Free Saline (CMF 10 x) NaCl (1.37 M) 80 g KCl (0.04 M) 3 g NaH2PO4 (0.004 M) 0.5 g KH2PO4 (2 M) 0.25 g NaHCO3 (0.12 M) 10 g Glucose (0.1 M) 20 g in 1000 ml distilled water, pH 7.3 9. β-galactosidase Detection Buffer 0.5 mg/ml 5-Bromo-4-Chloro-3-Indolylβ-D-galactopyranoside (X-gal) 25 mM potassium ferrocyanide 25 mM potassium ferricyanide 2 mM MgCl2 1 mM spermidine 0.02% Nonidet P-40 0.01% sodium deoxycholate in PBS 10. Chicken Embryo Fibroblast Media F10 medium 12.5% fetal calf serum 5% chicken serum 1.2% vitamin solution (100X) 1.2% folic acid 0.5% DMSO

6. Tris-NaCl Buffer 100 mM Tris-HCl, pH 7.5 150 mM NaCl

Section in Situ Hybridization The precise localization of gene expression within the tissue may not be immediately obvious from whole mount in situ hybridization. In situ hybridization on sections can identify whether a gene is expressed within the epithelium, the mesenchyme or both. Section in situ hybridization also alleviates possible probe penetration problems which sometimes occur with whole mount in situ hybridizations in larger embryos. Section in situ hybridization is performed as described by Nieto et al4 with some modification. Probes are labeled with digoxigenin using a digoxigenin RNA labeling kit (Boehringer Mannheim) and brought up in a standard volume of 100 µl. Briefly, paraffin sections are collected on fresh TESPA (30Aminopropyltriethoxysilan, Sigma) coated slides, dewaxed in xylene, taken through an ethanol series and postfixed in 4% paraformaldehyde for 30 minutes. Specimens are digested with proteinase K (10 µg/ml) for 5 minutes at 37°C, refixed with 4% paraformaldehyde and

398

Molecular Basis of Epithelial Appendage Morphogenesis

washed in Tris/glycine buffer (Table 20.1). Cultures are hybridized overnight at 60°C in section in situ hybridization buffer (Table 20.1) with a probe diluted 1:100. Posthybridization washes include 5 x SSC three times for 20 minutes followed by 0.5 x SSC in 20% formamide for 40 minutes at 60°C. The buffer is replaced with fresh buffer and allowed to cool to 37°C. Samples are then washed with STE (Table 20.1) for 15 min at 37°C, digested in STE containing 10 µg/ml RNase A, and rewashed in STE. The specimens are then washed in 0.5 x SSC in 20% formamide for 30 minutes at 60°C followed by 2 x SSC for 30 min at room temperature. The slides are preblocked with blocking solution (Table 20.1) and incubated with blocking solution containing alkaline phosphatase labeled sheep anti-digoxigenin Fab fragments (1:1500, Boehringer Mannheim) at room temperature overnight. Free antibody is removed by washing in Tris-NaCl (Table 20.1) followed by 0.1% Tween-20, 0.5 mg/ml levamisol. Antibody is detected with BM purple substrate in 0.1% Tween-20, 0.5 mg/ml levamisol. Color development at room temperature is stopped by the addition of a solution containing 10 mM Tris-HCl, pH 8.0, and 1 mM EDTA. The slides are mounted with glycerin mounting media.

Two Color in Situ Hybridization To analyze the order of appearance of two mRNA species or to see how two mRNAs are aligned, it is useful to use two color in situ hybridization. Two different transcripts can be detected on the same section using digoxygenin and fluorescein labeled riboprobes. Using the section in situ hybridization protocol outlined above, the mRNA with the strongest expression level is detected by hybridization to a fluorescein labeled probe which is detected with an anti-fluorescein antibody coupled to alkaline phosphatase using fast Red as a substrate. The mRNA with weaker expression levels is then hybridized to a digoxygenin labeled probe which is detected with an anti-digoxygenin antibody coupled again to alkaline phosphatase using BM purple (Boehringer Mannheim, Indianapolis, IN).

Whole Mount Immunohistochemistry To observe the three dimensional distribution of specific proteins, whole mount immunostaining is used. We use the procedure devised by Dent et al.5 Tissues are dissected in a dish containing PBS and fixed in 20% dimethylsulfoxide (DMSO)/80% methanol for 2 h at room temperature or at 4°C overnight. Following fixation, samples are bleached in 10% hydrogen peroxide (prepared from a 30% stock into 1:4 DMSO:MeOH) for at least 2 h (up to 4 days) to inactivate endogenous peroxidase. Samples are washed in TBST for 20 minutes and blocked with nonimmune serum for 30 minutes. The samples are then incubated with primary antibody in 95% fetal calf serum (FCS)/5% DMSO at room temperature overnight. After antibody incubation the tissues are washed for five 1 h washes in TBST. The tissue is then incubated with secondary antibody (biotinylated goat anti-rabbit or biotinylated horse anti-mouse IgG) diluted in 95% FCS/5% DMSO at room temperature overnight. Samples are washed five times for 1 h each wash in TBST. Tissues are incubated with tertiary antibody (ABC kit or Streptavidin-horseradish peroxidase (HRP), Vector Laboratories) diluted in 95% FCS/5% DMSO at room temperature for 2-3 h. The samples are washed three times for 30 min in TBST. Antibody staining is detected by incubating with an appropriate substrate (AEC or DAB, Vector Laboratories). Staining reactions are stopped by washing in TBS and the samples are mounted.

Section Immunohistochemistry A more precise localization can be obtained by immunofluorescence or immunoenzyme staining of a tissue section. By using double staining techniques the distribution of two

Current Methods in the Study of Avian Skin Appendages

399

molecules can be compared. These techniques have been used to demonstrate molecular expression in feather buds.6,7 Chick embryo or skin explants fixed in Bouin’s fixative for 1-2 h are washed overnight in 70% ethanol. Samples are then dehydrated through an ethanol series (80, 95, and 100%), cleared in xylene, and embedded in paraffin. Sections at 6-8 µm thickness are collected on gelatin-coated slides. The slides are deparaffinized and incubated with primary antibody (diluted in 1 x TBST containing 5% fetal calf serum and 1% bovine serum albumin (BSA)) overnight in a humidified chamber at room temperature. The sections are washed with TBST and incubated with alkaline phosphatase-conjugated secondary antibody followed by color development using NBT and BCIP as substrates (Promega).

Skin Explant Cultures Although in vivo experiments provide an excellent model to study the development of skin appendages, there are some drawbacks. It is difficult to do extensive manipulation, the whole embryo may dilute out the reagents, and 20-40% mortality is not unusual. Skin explant cultures are a useful method to complement in vivo experiments. Skin explants were originally started by Wolff and Haffen.8 Sengel9 has continued to develop the explant culture procedure and has stated that explants from E6.5 embryos still need fetal calf serum, but after E6.75 no serum is needed. The following is the procedure we use routinely.

Skin Explant Culture Pathogen-free chicken embryos are purchased from SPAFAS (Preston, CT). Eggs are incubated in a humidified incubator at 38°C and staged according to Hamburger and Hamilton.10 Stage 28-34 chicken embryo dorsal skin between the lower neck to the tail region are dissected in Hanks’s buffered saline solution (HBSS) and placed on culture insert in 6-well culture dishes (Falcon, Lincoln, NJ) containing 2 ml/well of Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 2% fetal calf serum and gentamicin (1:1000). Media is placed both in the outside well and the inner chamber. In the inner chamber, a thin layer of the medium is left to keep the explant moist and to provide an air-liquid interface. The skin explant cultures are incubated at 37°C at an atmosphere of 5% carbon dioxide and 95% air. The medium is changed every 2 days.

Epithelial-Mesenchymal Recombined Explants When epithelia and mesenchyma are separated, the organizations of both components are lost. However, if the epithelium and mesenchyme are recombined shortly after separation, feather germs regenerate (reviewed in refs. 9 and 11). If the epithelium is rotated from the mesenchymal orientation in the recombinants, the anterior-posterior orientation of the feather buds will follow that of the epithelium.12 This provides a unique opportunity to study the dependence of molecular expression on E-M contact and the order of molecular expression required for skin appendage formation. Stage 29-33 chicken embryo dorsal skins are dissected in HBSS and incubated in 2x calcium-magnesium free saline (Table 20.1) with 0.25% EDTA on ice for 10 minutes. The epithelium and mesenchyme are carefully separated using watchmaker’s forceps. The skin is then recombined with the epithelium and mesenchyme out of phase from their original positions in DMEM media with or without rotation to ensure that existing interactions occurring prior to the recombination are reset. The recombinants are placed on culture inserts in six-well culture dishes for growth in DMEM media with 10% fetal calf serum. Two milliliters of media are added to the culture dish and the skin is kept moist in the insert.

400

Molecular Basis of Epithelial Appendage Morphogenesis

Bead Mediated Local Delivery of Reagents to Skin Explant Localized delivery of growth factors utilizing Affi-Gel Blue beads (Bio-Rad, 100-250 µm in diameter) and Heparin-acrylic beads (Sigma, 200-250 µm diameter) are prepared as described in Hayamizu et al13 and Vogel and Tickle14 with some modifications. Both types of beads are washed twice in sterile PBS and approximately 100 beads are added to 5 µl aliquots of growth factor. The Affi-gel blue beads are incubated at 37°C for 1 h and the heparinacrylic beads are incubated at room temperature for 2 h before use. If treated beads are not immediately used, they are stored at 4°C for up to 1 week. Beads stored for longer than 1 week exhibit less activity in our skin explant culture assay. The treated beads are placed on top of the skin explants with fine tweezers. This method also can be applied to other reagents such as retinoic acid15 and cAMP homologue.16

Perturbing Functions with Antibodies, Growth Factors, and Drugs Cell adhesion is one of the key components of cell interaction, and cell adhesion molecules may play morphoregulatory roles. To explore the function of cell adhesion molecules during feather bud development, antibodies directed against different cell adhesion molecules can be added to the skin explant culture. Both polyclonal and monoclonal IgG or Fab fragments can be used. The antibodies are dialyzed against DMEM overnight and sterilized by filtering before adding to the culture medium.

Manipulation of Chicken Embryo Microsurgery and Microinjection For experimental manipulation in ovo, eggs are incubated lying on their side without shaking prior to infection for better access to the embryos. Eggs are moved to a tissue culture hood to maintain a sterile environment for surgical procedures. The eggs are sterilized with 70% ethanol, and about 2 ml of albumen are withdrawn with an 18 gauge needle to free the embryo from the egg shell. Care is taken not to damage the yolk. The surface of the egg shell is covered with tape and a 1 cm diameter circle is cut to expose the embryo. A little HBSS containing Gentamycin (1:100 dilution) is carefully dripped onto the blastoderm or embryo to keep it moist. India ink diluted 1:10 in PBS is injected beneath the blastoderm to render it clearly visible with a 1 ml syringe fitted with a 25 gauge needle. Care is taken to ensure that no air bubbles are present in the ink solution. The membrane is carefully penetrated and the needle advanced such that its tip comes to lie about 1 mm beneath the center of the blastoderm. India ink is gently injected until the entire embryo is visible above the ink background. For older embryos, ink injection is not necessary. Tungsten needles are used to remove targeted tissues for ablation studies. Host tissues to be used in grafting studies should be prepared in advance and stored in ice cold HBSS. Ablated tissues may be replaced with tissues from other regions of the embryo. After the manipulations, HBSS containing Gentamycin (1:100 dilution) is dripped onto the embryo to reduce the risk of contamination. All membranes should be replaced over the embryo. Microcapillary needles are filled with 3-6 µl of reagents for microinjection. The reagents (viral vector, dye, drugs) are gently injected into the dorsal body ectoderm of stage 10-20 embryos or other sites of interest. A microinjection apparatus is useful when the injection site is small. For older embryos, a syringe converted injector, with connection to polyethylene tubing and capillary tubing, can work. After injection, a little HBSS is added to the embryo and the window is carefully covered with scotch tape. The egg is placed in a humidified incubator at 38°C. Embryos are harvested at stages 35-38 and analyzed for transgene expression, the distribution of dyes and changes in skin morphology.

Current Methods in the Study of Avian Skin Appendages

401

Delivery of Reagents to Eggs If systematic delivery is the goal, reagents can be delivered easily through the chorioallantoic veins. Eggs are incubated horizontally from E1. On the day of surgery, eggs are prepared as described above (for eggs older than E7 or so, it is not necessary to withdraw albumin). A small window is made and a drop of media is dripped to wet the chorioallantoic membrane. Fiberoptic light in a darkened room can help to localize the embryos. Reagents such as retinoic acid are then injected into the sac around the embryo. The window is then sealed by scotch tape.

Chorioallantoic Membrane Cultures The chicken embryo skin explant can usually be cultured in the insert dish for 5-7 days; then they stop elongating due to a lack of nutritional penetration. In order to keep the skin explant growing, we have cultured the explant on a chicken embryo chorioallantoic membrane.17 The recipient eggs are incubated vertically for 8 days and moved to a tissue culture hood. The eggs are sterilized with 70% ethanol and a window (1 cm in diameter) is cut on top of the eggs to expose the embryo. The egg shell and underlying membrane is carefully peeled back using tweezers. The cultured skin is transferred with its dorsal face up, to the top of the chorioallantoic membrane. For better skin adhesion to the membrane, the chorioallantoic membrane can be scratched with tweezers so the capillary vessels bleed a little. Care must be taken not to damage the larger vessels; otherwise the embryos will die because of the bleeding. A little HBSS with antibiotic is added to wet the skin and the chorioallantoic membrane. The window is then sealed with scotch tape and the egg is kept at 38°C without shaking. The explant should be checked every day and moistened with HBSS. Usually after 2-3 days of culture, blood vessels can be seen in the feathers.

Monitoring Cell Behavior Morphogenesis is based on cell behaviors. When morphogenesis occurs, it is relevant to know whether the change is due to alteration in cell proliferation, migration or death. The following procedures are helpful for these analyses.

Retroviral Lineage Tracing Replication defective retroviral vectors offer the ability to trace lineage over an extended period of time, since the virus is only passed on to daughter cells and is not diluted in subsequent generations.18,19 Plasmids encoding replication defective retroviral vectors are transfected into packaging cell lines by lipofection (GIBCO). Transfected cells expressing the viral Neor marker are selected for G418 resistance. The media containing retrovirus are filter sterilized, concentrated by centrifugation and stored as frozen stock. Viral titers can be assessed by transducing susceptible cells with different viral dilutions. By microinjecting the retrovirus to target areas, the area of infection can be limited.20 The methods used to detect the viral transduction are vector-dependent. Retroviruses expressing β-galactosidase are detected by X-gal histochemistry. For this procedure, tissues are incubated in β-galactosidase detection buffer (Table 20.1) until a blue color develops. The lineage and migration of living cells expressing β-galactosidase can be traced using the Imagene fluorescent substrate (Molecular Probes).21 For this procedure, skins are immersed in 20 µM ImaGene substrate for 30 min, rinsed and viewed with fluorescence microscopy using a SIT-66 videocamera (Dage MTI, Michigan City, IN). Unfortunately, the precise location of the initially transfected cells cannot be ascertained, which is a disadvantage of retroviral lineage tracing.

402

Molecular Basis of Epithelial Appendage Morphogenesis

Fluorescent Dye Tracing Cell migration and lineage can also be examined by injecting the fluorescent, lipophilic, carbocyanine dyes, Di A, Di I, Di O or their derivatives. These dyes integrate into cell membranes (Molecular Probes).22 This procedure is frequently used in developmental neurobiology, and can be applied to skin appendage research. These nontoxic dyes offer several advantages: The precise identity of the labeled cells can be established, the dyes do not spread to neighboring cells and are therefore confined to labeled cells and descendants, they can be detected in living tissues, fixed whole mount samples or histological sections. Neighboring lineages can be distinguished from one another within an individual organism by using different labeling moieties. Unfortunately, the dye is diluted by mitotic cell divisions and can only be followed for a limited time period.

Cell Proliferation for Detection in Sections Cell proliferation also plays a major role in the formation of histological structures. Proliferating cells can be detected in tissues by the incorporation of [3H] thymidine or bromodeoxyuridine (BrdU). Skin explants are labeled by adding [3H] thymidine (10 µCi/ml) to the in vitro culture media. The samples then are fixed, sectioned and dipped in NTB2 emulsion (Kodak) for autoradiography. Staining with BrdU can localize the staining more precisely. For this procedure, tissues are incubated with BrdU for 1h, fixed in 2% paraformaldehyde, dehydrated and sectioned. Samples are then washed with PBS. The cellular DNA is denatured in 2 M HCl for 20 min and neutralized with PBS for 5 min. The BrdU labeled DNA is stained with monoclonal anti-BrdU antibody (Boehringer Mannheim), diluted 1:25 for 30 min, followed by biotinylated horse anti-mouse antibody, and then detected with the ABC and AEC kits (Vector Laboratories).

Cell Proliferation for Whole Mount Detection Labeling with BrdU offers the ability to view the proliferating cells using whole mount staining. For whole mount, tissues are dissected in a dish containing serum-free DMEM, pulse labeled with 20 µM BrdU (diluted 5mM BrdU 1:100 in DMEM without serum) in a 37°C incubator for the desired length of time and fixed by immersion in absolute methanol at room temperature for 2-3 h or at 4°C overnight. The samples are then bleached with 10% hydrogen peroxide in 1:4 DMSO:100% MeOH for 2 h to overnight and rehydrated in four 15 minute PBS washes. DNA is denatured by incubating the tissues in 2N HCl at 37°C for 60 minutes followed by neutralization through immersion in four 10 minute 0.1 M sodium borate buffer, pH 8.5 washes and four 10 minute washes in PBS. The samples are incubated with enough anti-bromodeoxyuridine antibody (approximately 3 µg/ml antibody diluted in PBS with 0.1% BSA) to cover the tissue overnight at 4°C. Samples are then washed 5 times in PBS for 20 minutes and incubated with enough secondary antibody (biotinylated horse anti-mouse IgG) for 3 h at room temperature at 4°C. The antibody is diluted in PBS containing 0.1% BSA. Samples are again washed 5 times in PBS for 20. Samples are incubated with the tertiary antibodies (we use the ABC kit from Vector Laboratories) for 2-3 h and then washed 5 times in PBS for 20 minutes. BrdU staining is detected by adding the AEC substrate and observing closely for color development. The color reaction is stopped by washing with PBS 3 times. Samples are cleared in 50% glycerol (in PBS) overnight and then mounted in 80% glycerol.

Tracing Apoptosis Programmed cell death plays an important role in a number of tissues. In the skin appendages, it functions to form the space between feather barbs and is also involved in the hair cycle. Apoptosis produces DNA fragmentation which can be determined by in situ

Current Methods in the Study of Avian Skin Appendages

403

end-labeling.23,24 For this procedure, paraffin tissues are washed in xylene to remove the paraffin, rehydrated and protease treated. The tissues are then end-labeled with deoxynucleotides containing biotin-11-dUTP or digoxigenin-11-dUTP. The labeled DNA is detected with avidin or anti-digoxigenin antibodies. Labeled apoptotic cells can be distinguished from necrotic cells by microscopic examination. In apoptotic cells, DNA fragmentation occurs hours prior to cell death. Therefore, apoptotic cells can also be distinguished from necrotic cells by photolabeling with ethidium monoazide, which is excluded by living cells but intercalates into the DNA of necrotic cells.

Perturbing Functions with Retroviral Mediated Gene Transduction Retrovirus Mediated Gene Delivery Retroviral mediated gene delivery is an efficient method to place exogenous genes into cells of an organized tissue. The retroviruses are a large family of RNA viruses, primarily infecting vertebrates (reviewed in ref. 25). The retroviral life cycle has several unique features that make it useful as a vector for gene delivery to the cell. Retroviruses must bind to specific cell surface receptors in order to penetrate the cell. Once in the cytoplasm, the viral RNA is reverse transcribed into a DNA intermediate that becomes stably integrated into the host cellular genome. The virus then uses the host cellular transcription machinery to synthesize new viral mRNA which is assembled into infectious virions to infect adjoining cells. This is a highly efficient process. In some cases, the viral transcripts can account for 10 per cent of the total mRNA transcribed by the infected cell.26 The retroviral life cycle does not necessarily harm the infected cell. In most cases, retroviral transduction merely produces a cell which is permanently capable of producing infectious virions at a low level. The integrated viral genome is also vertically transmitted to daughter cells, ensuring the stable transmission of exogenous genes. These features of stable gene expression and minimal host toxicity have made retroviruses attractive as vectors for gene delivery in a number of culture systems.

RCAS Retrovirus Exogenous genes can be efficiently introduced to cells within a developing tissue using a replication competent retrovirus. The RCAS BP retroviral vector (Replication Competent Avian Sarcoma virus LTR, with a splice acceptor and sequences derived from the polymerase gene of the Bryan high titer Rous sarcoma virus) has been developed by the Hughes laboratory.27-29 The RCAS BP was derived from the Rous sarcoma virus (RSV), a member of the ASLV (avian sarcoma and leukemia virus) family. The RSV virus contains a 2 kb src oncogene sequence plus the usual viral proteins, gag, pol and env. Hughes and colleagues excised the src gene and replaced it with a Cla I restriction site for subcloning exogenous transgenes.

Vector Construction Plasmids containing the genes of interest are excised with appropriate restriction endonucleases and placed into the multicloning site of the pClaNcoHa vector.30 This vector adds a hemagglutinin tag to the transduced protein for easy identification and purification. The fragment of interest is isolated by digestion with Cla I and subcloned into the Cla I cloning site of the RCAS BP retroviral vector.28 The orientation of the vector is determined by restriction endonuclease digestion using a site within the subcloned fragment and one within the vector. RCAS BP can stably integrate up to 2.4 kb of insert.31 To date, the RCAS retrovirus has been used most extensively to introduce foreign genes to avian tissues. However, other retroviruses (e.g., spleen necrosis virus) and other viruses

404

Molecular Basis of Epithelial Appendage Morphogenesis

(adenovirus) are also used. A more comprehensive description of techniques involving the use of RCAS is available.31

Viral Production Dorsal skin fibroblasts from pathogen-free SPAFAS chicken embryos are used as retrovirus producing cells. Cells plated at 1 x 106 cells per ml in 35 mm dishes (Falcon) are cultured in DMEM with 10% fetal calf serum. At 24 h, cells are transfected with 1 µg of RCAS BP DNA containing the gene of interest using 10 µl of lipofectamine (Life Technologies) in 1 ml of DMEM. DMEM, and 20% FCS are added at 4 h to terminate the transfection reaction. 24 h after transfection, the media is changed to chicken embryo fibroblast media (Table 20.1). The cells are grown until they reach 70% confluence. They are then transferred to 100 mm dishes (Falcon). When the cells again reach 70% confluence, fresh complete avian media is added and the retrovirus containing media is collected after 24 h. Viral containing media is filtered through a 0.45 µm surfactant-free cellulose acetate filter (Nalgene) and stored in aliquots at -70°C. The viral titer is obtained as described by Morgan and Fekete.31 A number of chicken lines with different resistance or susceptibility to viral infection have been described.31 Among these, Line 0 and standard pathogen-free eggs from SPAFAS can be infected by the A-subgroup retroviruses, but Line 72 is resistant to the A-subgroup retroviruses. Line 15b1 can be infected by the E-subgroup retroviruses, but Line 0 is resistant to the E-subgroup retroviruses. Lines 0, 72 and 15b1 are available from the USDA Poultry Research Laboratory in East Lansing, MI.

In Search of Candidate Genes To study the molecular cascade involved, we need to identify the involved genes and not be prejudiced that some known “important” genes can do everything. How do we identify the candidate genes? One of the most powerful approaches in molecular analysis is through the isolation of mutants and identification of genes involved in that particular pathway. This has been very successful in the study of Drosophila. It also has been powerful in the study of mouse mutants and human genetic diseases. Unfortunately, in avians, in addition to the macro-chromosomes there are a large number of microchromosomes that are nearly undistinguishable.32 Thus, knowledge about the avian genome is far behind mouse and human genome. This makes it difficult to analyze avian mutants or to isolate genes even when mutants are available. This is the reason that we have to rely more heavily on the “borrowed genetics” approach (see below). However because the integument is easy to identify, there are indeed mutants involving plumage, comb, and scales.33 Among them, only scaleless34 has been studied in much detail.35,36 It is an autosomal recessive mutation; the primary defect was shown to be in the epithelium using epithelial-mesenchymal recombination and explant culture.37 Recently, scaleless skin was shown to have abnormal Shh expression.38 The scaleless phenotype can be partially rescued by exposure to exogenous FGF.39 However, we still do not have a direct handle on the scaleless gene. Another approach is to compare the difference of gene expression at different stages (e.g., before and after induction) of appendage morphogenesis or between wild type and mutants using biochemical, immunological or molecular biological methods. This can be pursued using two dimension gel analysis or monoclonal antibody screening to identify the difference in protein expression profiles. It can also be pursued by studying the difference in transcripts using differential display,40 or by subtraction library analyses. It is possible to identify some molecules that are differentially expressed. These candidate molecules can then be tested using the flowchart in Figure 20.1. The pitfall is that many candidates may be identified, and it is difficult to know which one is the causal one.

Current Methods in the Study of Avian Skin Appendages

405

Table 20.2. Sources for research relating to chicken embryos Avian Disease and Oncology Laboratory 3606 E. Mount Hope Rd. East Lansing, Michigan 48823 Phone (517) 337-6828 Suppliers of specific lines of viral resistant and viral susceptible chicken embryos. SPAFAS Connecticut Hatchery 190 Route 165 Preston, CT 06365-8531 Phone (860) 889-1389 FAX (860) 889-1991 Suppliers of pathogen-free viral susceptible chicken embryos. They are from the original Mt. Hope Leghorn. Genetic Resources Conservation Program University of California, Davis Davis, CA 95616 Phone (916) 754-8501 FAX (916) 754-8505 e-mail [email protected] Suppliers of genetic mutant stocks of chicken embryos University of Connecticut at Storrs Department of Poultry Genetics White Building, Room 9 3636 Horsebarn Rd Ext Storrs, CT 06269-4039 Phone (860) 486-1016 Suppliers of genetic mutant stocks of chicken embryos Poultry Genetic Stock Listing with Lists of Breeders and Suppliers International Registry of Poultry Genetic Stocks, RG Somes, Storrs Agricultural Experiment Station, University of Connecticut, Storrs, CT World Wide Web Sites for Chicken Genome Maps US Poultry Gene Mapping Michigan State University East Lansing, Michigan http://poultry.mph.msu.edu/ ChickMap Roslin Institute Edinburgh, United Kingdom http://www.ri.bbsrc.ac.uk/chickmap/ChickMapHomePage.html

“Borrowed Genetics” Approach Many new genes have recently been cloned in mutants of other species. The phenotypes imply that they are functionally involved. We and others have found that it is beneficial to clone chicken homologs of genes known to be involved in development in other known genetic models. For preliminary tests, we first determine the molecular distribution of the gene of interest to see whether the pattern is interesting to feather development. For this purpose, we need to isolate in situ hybridization probes from chicken cDNAs. To clone

406

Molecular Basis of Epithelial Appendage Morphogenesis

genes identified in other species, we use PCR or homology screening. If the gene has been identified in only one species, we use homology screening: if it has been cloned in several species other than the chicken, we use PCR based on regions with conserved amino acid sequence.

PCR Screening mRNA from appropriately aged embryos is prepared using Trizol Reagent (Life Technologies) and cDNA is reverse transcribed using oligo-dT priming and AMV reverse transcriptase. A pair of degenerate primers is designed based on a comparison of the conserved amino acid sequences, trying to minimize the degree of degeneracy. We generally use primers greater than 17 nucleotides in length with a maximum degeneracy of 1024. Conditions for the PCR reactions are determined based on the predicted melting temperatures of the degenerate primers. Amplification reactions are performed with the pfu DNA polymerase (Stratagene). An aliquot of the PCR products are run on 1% agarose gels to verify that they are the expected size. They are subcloned by blunt-end ligation into PCR-Script (Stratagene). This vector offers Lac Z blue/white selection for vectors containing inserts when colonies are plated on media containing X-gal and IPTG. White colonies containing inserts are screened by miniprep DNA purification.41 The identity and orientation of the PCR products are verified by dideoxynucleotide sequencing using Sequenase 2 (USB) from the available T3 and T7 promoter primers within the PCR-Script vector. These constructs can then be used to generate in situ hybridization probes.

Homology Screening We have prepared a stage 29-34 chicken embryo cDNA library in the Lambda Zap II vector kit (Stratagene) following the company’s recommended procedure. The phagemid library is grown at a density of 50,000 plaques per plate on XL-1 blue mrf- cells in 150 mm dishes and transferred to nitrocellulose in duplicate for hybridization screening. Probes, based on PCR fragments or genes from other species, are radiolabeled by random priming using standard procedures and added to the hybridization buffer overnight at an appropriate temperature for the specific probes. Free and nonspecifically bound probe are removed by washing in progressively higher stringency conditions (lower salt and higher temperature). The specificity of binding while washing can be monitored with a Geiger counter. Autoradiographs from the duplicate hybridized filters are aligned. Positive plaques should be found on both filters to warrant further screening. The agar plate containing the positive plaques is removed with the large end of a sterile Pasteur pipet, diluted in SM buffer and replated. We find that 2-3 rounds of hybridization screening are required to plaque purify the phage of interest. Clones of interest are rescued from the phagemid by in vivo excision following the company’s recommended procedure.

Perspectives In summary, with the rapid development of molecular biology in the last two decades, the technology is now available to address issues concerning the molecular basis of skin appendage morphogenesis. The classical embryology experiments (reviewed in ref. 9) serve as a reservoir of fascinating phenomena. With the methods described here, we hope the new generation of skin appendage research scientists will advance this field to new heights.

Acknowledgment We thank Dr. Jackie Pisenti for suggestions on information about chicken mutants. This work is supported by grants from NIH and NSF.

Current Methods in the Study of Avian Skin Appendages

407

References 1. Dieffenbach CW, Dveksler G. PCR primer. New York, Cold Spring Harbor Laboratory Press, 1995. 2. Sasaki H, and Hogan BL. Differential expression of multiple fork head related genes during gastrulation and axial pattern formation in the mouse embryo. Development 1993; 118:47-59. 3. Wilkinson DG. Whole mount in situ hybridisation of vertebrate embryos. Oxford, UK: Oxford University Press, 1992. 4. Nieto MA, Patel K, Wilkinson DG. In situ hybridization analysis of chick embryos in whole mount and tissue sections. Meth Cell Biol 1996; 51:219-235. 5. Dent JA, Polson AG, Klymkowsky MW. A whole-mount immunocytochemical analysis of the expression of the intermediate filament protein vimentin in Xenopus. Development 1989; 105:61-74. 6. Chuong CM, Edelman GM. Expression of cell-adhesion molecules in embryonic induction. I. Morphogenesis of nestling feathers. J Cell Biol 1985; 101:1009-1026. 7. Chuong C-M, Oliver G, Ting S et al. Gradient of homeoproteins in developing feather buds. Development 1990; 110:1021-1030. 8. Wolff E, Haffen K. Sur une méthode de culture d’organes embryonnaires in vitro. Texas Rep Biol Med 1952; 10:463-472. 9. Sengel P. Morphogenesis of skin. In: Abercrombie M, Newth DR, Torrey JG, eds. Cambridge: Cambridge University Press, 1976. 10. Hamburger V, Hamilton HL. A series of normal stages in development of the chick embryo. J Morphol 1951; 88:49-91. 11. Sengel P. Feather pattern development. In: Ciba Foundation Symposium 29. New York: Elsevier Science, 1978:51-70. 12. Novel G. Feather pattern stability and reorganization in cultured skin. J Embryol Exp Morph 1973; 30:605-633. 13. Hayamizu TF, Sessions SK, Wanek N et al. Effects of localized application of transforming growth factor beta 1 on developing chick limbs. Dev Biol 1991; 145:164-173. 14. Vogel A, Tickle C. FGF-4 maintains polarizing activity of posterior limb bud cells in vivo and in vitro. Development 1993; 119:199-206. 15. Chuong C, Ting SA, Widelitz, RB et al. Mechanism of skin morphogenesis: II. Retinoic acid modulates axis orientation and phenotypes of skin appendages. Development 1992; 115:839-852. 16. Noveen A, Jiang T-X, Chuong C-M. Protein kinase A and protein kinase C modulators have reciprocal effects on mesenchymal condensation during skin appendage morphogenesis. Dev Biol 1995; 171:677-683. 17. Rawles, M.E. Tissue interactions in scale and feather development as studied in dermalepidermal recombination. J Embryol exp Morph 1963; 11:765-789. 18. Fekete DM, Cepko CL. Retroviral infection coupled with tissue transplantation limits gene transfer in the chicken embryo. Proc Natl Acad Sci USA 1993; 90:2350-2354. 19. Galileo DS, Gray GE, Owens GC et al. Neurons and glia arise from a common progenitor in chicken optic tectum: demonstration with two retroviruses and cell type-specific antibodies. Proc Natl Acad Sci, USA 1990; 87:458-462. 20. Widelitz RB, Chuong C-M. Replication-defective virus infection of feather buds produces a localized region of β -galactosidase activity. Biochem Biophys Res Commun 1992; 186:1020-1024. 21. Westerfield M, Wegner J, Jegalian BG et al. Specific activation of mammalian Hox promoters in mosaic transgenic zebrafish. Genes and Dev 1992; 6:591-598. 22. Honig MG, Hume RI. DiI and DiO: Versatile fluorescent dyes for neuronal labeling and pathway tracing. Trends Neurosci 1989; 12:333-336. 23. Wijsman JH, Jonker RR, Keijzer R et al. A new method to detect apoptosis in paraffin sections: In situ end-labeling of fragmented DNA. J Histochem Cytochem 1993; 41:7-12.

408

Molecular Basis of Epithelial Appendage Morphogenesis

24. Gold R, Schmied M, Rothe G et al. Detection of DNA fragmentation in apoptosis: Application of in situ nick translation to cell culture systems and tissue sections. J Histochem Cytochem 1993; 41:1023-1030. 25. Miller AD. Retroviral vectors. Curr Topics Microbiol Immunol 1992; 158:1-24. 26. Coffin JM. Retroviridae and their replication. In: Fields BN, Knipe DM, eds. Fundamental Virology. Philadelphia, Pa: Raven Press Ltd., 1991. 27. Hughes S, Kosik E. Mutagenesis of the region between env and src of the SR-A strain of Rous sarcoma virus for the purpose of constructing helper-independent vectors. Virology 1984; 136:89-99. 28. Hughes SH, Greenhouse JJ, Petropoulos CJ et al. Adaptor plasmids simplify the insertion of foreign DNA into helper-independent retroviral vectors. J Virol 1987; 61:3004-3012. 29. Petropoulos CJ, Hughes SH. Replication-competent retrovirus vectors for the transfer and expression of gene cassettes in avian cells. J Virol 1991; 65:3728-3737. 30. Laufer E, Nelson CE, Johnson RL et al. Sonic hedgehog and FGF-4 act through a signaling cascade and feedback loop to integrate growth and patterning of the developing limb bud., Cell 1994; 79:993-1003. 31. Morgan BA, Fekete DM. Manipulating gene expression with replication-competent retroviruses. Methods Cell Biol 1996; 51:185-218. 32. Bitgood JJ, Shoffner RN. Cytology and cytogenetics. In: Crawford RD, ed. Poultry Breeding and Genetics. Amsterdam: Elsevier, 1990:401-428. 33. Somes RG Jr. Mutations and major variants of plumage and skin in chickens. In: Crawford RD, ed. Poultry Breeding and Genetics. Amsterdam: Elsevier, 1990:169-208. 34. Abott UK, Asmundson VS. Scaleless, an inheritied ectodermal defect in the domestic fowl. J Hered 1957; 48:63-70. 35. Brotman HF. Abnormal morphogenesis of feather structure and pattern in the chick embryo integument. I. Macroscopic description. J Exp Zool 1976; 196:323-340. 36. Brotman HF. Epidermal-dermal tissue interactions between mutant and normal embryonic back skin: Site of mutant gene activity determining abnormal feathering is in the epidermis. J Exp Zool 1977; 200:243-257. 37. McAleese SR, Sawyer RH. Correcting the phenotype of the epidermis from chick embryos homozygous for the gene scaleless (sc/sc). Science 1981; 214:1033-1034. 38. Ting-Berreth SA, Chuong C-M. Sonic hedgehog in feather morphogenesis: Induction of mesenchymal condensation and association with cell death. Dev Dyn 1996; 207:157-170. 39. Song H, Wang Y, Goetinck PF. Fibroblast growth factor 2 can replace ectodermal signaling for feather development. Proc Natl Acad Sci USA 1996; 93:10246-10249. 40. Liang P, Pardee AB. Differential display of eukaryotic messenger RNA by means of the polymerase chain reaction. Science 1992; 257:967-971. 41. Sambrook J, Fritsch EF, Maniatis T. Molecular cloning: A laboratory manual. New York, Cold Spring Harbor Laboratory Press, 1989.

CHAPTER 21

Molecular Biology of Anhydrotic Ectodermal Dysplasia Juha Kere

Introduction

H

istorically, the phenotype of anhydrotic ectodermal dysplasia (EDA) has stimulated considerable interest among investigators. A famous early description of the disorder was written by Charles Darwin1 (cited in more easily accessible sources2,3), even though an earlier report4 exists. In his description, Darwin also accurately listed all hallmarks of the X-linked pattern of inheritance that later defined the chromosomal localization of the gene. During the first half of this century, tens of cases were recorded in the literature.5-12 Other names for the syndrome have also been proposed, such as Christ-Siemens-Touraine (CST) syndrome and hypohydrotic ectodermal dysplasia (HED), because in most patients the syndrome manifests as hypohydrotic (i.e., reduced sweating) rather than anhydrotic (absent sweating).13 The present author prefers the older name “anhydrotic ectodermal dysplasia” because the acronym HED may be confused with a different syndrome, hydrotic ectodermal dysplasia (or Clouston syndrome) that is most often inherited as an autosomal dominant trait.14 Thus, genetically and biochemically, hydrotic ectodermal dysplasia and EDA are definitely distinct. In addition, more than 150 other ectodermal dysplasias have been delineated through clinical classification.14,15 Since no genes other than EDA have been identified and nothing is known of the biochemical basis in any of the other ectodermal dysplasias, the syndromic classification is commonly used in clinical diagnostics and genetic counseling. EDA, besides being the classical type of ectodermal dysplasia, is also by far the most common of them.16 This chapter discusses the phenotype of EDA, the best-characterized homologous animal model, the Tabby mouse, genetic mapping and cloning of the EDA gene, and finally, current clues to the function of EDA in ectodermal development.

The Phenotype of Anhydrotic Ectodermal Dysplasia and Carrier Manifestations The classical triad of EDA includes absent or reduced sweating (anhydrosis or hypohydrosis), missing or sparse hair (atrichosis or hypotrichosis), and missing or misshaped teeth (anodontia or hypodontia), combined with an X-linked recessive mode of inheritance (Fig. 21.1).1,4-15 Occasionally, nails may be abnormal, but this is much more typical of the dominantly inherited hydrotic ectodermal dysplasia.14,17 A consistent feature of the syndrome is a characteristic facies, with prominent forehead and supraorbital ridges, low nasal bridge and flat nose, deep set eyes and dry, wrinkled, and Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

410

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 21.1. Phenotypes of anhydrotic ectodermal dysplasia. The triad of signs typical of anhydrotic ectodermal dysplasia includes absent or malformed teeth (left two panels), thin and sparse hair (right panel), and lack of sweating.

hyperpigmented periorbital skin, prominent cheek bones, thick and everted lips, and small chin. Skin and subcutaneous fat are thin in all parts of the body. Atopy, dryness of the mucous membranes, eyes and airways, and recurrent bronchitis are common symptoms. Other, occasionally occurring features include the absence of nipples, hyperkeratosis of palms and soles, cleft lip and palate, and neonatal peeling of the skin.17-20 The number of missing teeth varies. Very rarely there is a complete lack of teeth or anodontia, but more commonly the remaining teeth are small and their shapes are aberrant: They are commonly peg-shaped and the number of cusps is reduced. Lack of sweat glands may lead to hyperthermia in hot weather and during physical exercise, even causing convulsions, subnormal intelligence (attributed to recurrent, severe hyperthermic periods during infancy and childhood), or death. Hair is usually fair, thin, and sparse, and eyebrows and eyelashes are absent. Axillary and pubic hair as well as beard may grow more strongly. Skin histology reveals that eccrine sweat glands are few, rudimentary or missing altogether, whereas apocrine glands may be normal.17,20 Female carriers of the syndrome often show incomplete symptoms and signs: They may have missing or abnormal teeth, their hair may be thin, and they may suffer from hot weather.21 Sweat glands can be counted and their function studied by staining skin with starch and iodine or other dyes that change color when hydrated by sweat. Such studies of carrier females reveal striped or patched patterns, consistent with the lines of Blaschko.22,23 The clinical signs are, however, unreliable in determining the carriership status of female relatives of EDA patients. Taken together, these clinical features and observations suggest the first clues to the molecular role of the EDA gene product: 1. EDA must be involved in a pathway common to ectodermal development in skin, teeth, and perhaps other tissues (cranial dysmorphology); 2. it must exert its function during fetal development, consistent with the timing of skin appendage and tooth morphogenesis; 3. it may continue to have a function in adult tissues (occasionally abnormal nails, thin skin); and 4. it must act locally in the skin rather than act humorally or diffusely over long distances (local effects in carrier females). Based on the clinical features it is not possible to infer which cell types in developing skin or teeth would express EDA protein. As reviewed in detail elsewhere in this book, ectodermal morphogenesis involves epithelial-mesenchymal interactions and, in principle, a defect in either of these tissues would be consistent with all phenotypic features.24

Molecular Biology of Anhydrotic Ectodermal Dysplasia

411

Phenotypic Similarity of the Tabby Mouse and Clues for Site of Action Falconer25 described Tabby (Ta), “a totally sex-linked gene in the house mouse” whose phenotype was characterized by abnormalities of hair, teeth, and some exocrine glands. These features are all present also in mice with the autosomal recessive mutations downless (dl) and crinkled (cr), and the dominant mutation sleek (Slk).26 Studies of tooth abnormalities, sweat gland morphology, and sex linkage in both mouse and human led to the proposal that EDA and Ta might be homologous genes in the two species.27 The distribution of hair and skin glands is different in mouse and human, so that mice have sweat glands only in their paw pads. In Ta mice, these structures are missing and no sweat production can be detected.27,28 Detailed molecular mapping studies have confirmed the syntenically corresponding positions of the EDA and Ta genes.29 Recombination grafting of epidermal and dermal layers from wild-type and Ta mice has been used to study the effect of the Ta mutation.30-32 A study of recombined tail skin sections cultured on hen’s egg tissue resulted in only partial loss of hair follicle formation and failed to assign the gene’s function in either skin layer.30 However, a different assay system, with recombined body skin sections grown in mouse testes, and hair morphology as the measure of mutation effect, assigned the function of Ta unequivocally to epidermis.31 A third study32 addressed both aspects by recombining both tail and body skin sections, growing them on nude mice, and observing both follicle formation and hair type. The results confirmed those of both previous studies: The function of Ta in tail follicle initiation could not be assigned to either epidermis or dermis, whereas body hair morphology was influenced solely by the type of epidermis. The authors speculate that this difference may be due to the different timing of developmental events in tail and body skin, and resulting different biochemical environments or changing expression patterns of the gene itself.32 Taken together, these results suggest that the Tabby gene might have more than just one role, perhaps in both epithelial and mesenchymal cell types. The first clues of the biochemical pathway with which Ta (and EDA) might be involved came from a series of observations on the role of epidermal growth factor (EGF). The effect of the Ta mutation was observed to be opposite to that of EGF, and EGF injections induced sweat glands and reversed the delayed eyelid opening and incisor eruption in newborn Ta mice.33 A more detailed study of the EGF pathway in both Ta mice and EDA patients indicated that EGF binding was reduced in EDA and Ta fibroblasts, with reduced EGF receptor (EGFR) expression at both protein and mRNA levels.34 Knock-out of EGFR causes epithelial immaturity and disorganization which is, however, strain-dependent, suggesting distinct modifying effects by other genes.35-37 The mechanism by which EDA and Ta affect EGFR expression, and whether the effects of EDA and Ta are mediated only through the EGF pathway, remain subjects of future studies.

Genetic and Physical Mapping of EDA and TA Even though EDA is an X linked recessive disease, occasional female patients have been observed who manifest the full phenotypic features of EDA. There are three general possibilities to explain these cases, namely: 1. they might be phenocopies, possibly caused by mutations in another gene that is involved in the same molecular pathway as the EDA gene; 2. they might be carriers of a single copy of a mutated EDA gene, but have an unusual inactivation pattern of X chromosomes, leading to an unusually severe expression of the phenotype; or 3. they might carry defective genes in both of their X chromosomes, leaving them without any functional copy.

412

Molecular Basis of Epithelial Appendage Morphogenesis

The diagnosis of a chromosomal translocation t(X;9)(q12;p24) in a female dubbed “AnLy” led EDA to be the first X chromosomal gene whose position along the chromosome was deduced.2,38,39 More recently, two additional translocations in female patients have been published, one with t(X;12)(q13.1;q24) and the other, dubbed “AK”, with t(X;1)(q13.1;p36).40,41 The reason for the full EDA phenotype in these patients is nonrandom X inactivation caused by X;autosome translocations. The translocated X chromosome remains active in all cells whereas the normal X is late replicating and inactive.40-42 Thus, the presumably normal copy of the EDA gene in the structurally normal X chromosome becomes nonfunctional and, because the translocation is associated with the clinical phenotype, the translocation must disrupt the EDA gene. Genetic linkage studies were undertaken by several groups to confirm and refine the genetic localization of the EDA gene.43-47 These studies also allowed predictive testing and carrier diagnosis in informative families.47 Important observations were made concerning genetic homogeneity of EDA: While some families may have been unlinked to the locus in Xq13, evidence suggested that most (95%) of the families included in large series were linked to the same locus.48 In order to clone and identify the EDA and Ta genes, physical mapping of the corresponding regions in human and mouse chromosomes was necessary. Comparative mapping in the mouse placed Ta between the androgen receptor (Ar) and Ccg1 genes, fully consistent with the human gene map.29 This result, and the finding of cross-hybridizing mouse probes that identified homologous segments in other species, including human, further supported the inference of evolutionary conservation between the EDA and Ta genes.29 In human, cloned markers and preliminary physical maps based on rare-cutter restriction analysis of genomic DNA were initially used,48,49 but a more comprehensive physical map was obtained when the whole region was first isolated in yeast artificial chromosome (YAC) clones.50 Physical maps and further probes allowed the fine mapping of translocation breakpoints and the recognition of male patients with submicroscopic deletions.50-53 Disappointingly, the vicinity of the X chromosomal translocation breakpoint in AnLy did not seem to contain coding DNA sequences, and a gene included in the locus DXS732 (isolated based on homology with a mouse CpG island29) mapped 200 kb centromeric of the AnLy breakpoint. However, these studies provided the necessary DNA clones and physical landmarks to proceed to identify the EDA gene.

Cloning of the EDA Gene and Structure of a Predicted Protein The systematic study of CpG islands isolated near the translocation breakpoints led to a startling observation that subsequently allowed the cloning of the EDA gene. One CpG island, unsurprisingly associated with a transcript on Northern blots, also detected a molecular rearrangement in DNA from patient AK, consistent with the t(X;1) breakpoint.54 Sequencing of the CpG island clones and identification of a cDNA in a sweat gland library verified the existence of a gene that turned out to span 200 kb of genomic DNA and consist of two exons.3 Even though the cloned cDNA was complete with a polyadenylation signal sequence and poly(A) tail, it was obvious that parts, most likely extending further 3' of exon 2, had not yet been sequenced. Northern analyses suggested the presence of alternatively spliced forms of the gene in several tissues, or possibly the existence of homologous genes elsewhere in the genome.3 Very recently, we have been able to identify an additional exon and alternative transcript (S. Ezer et al, work in progress). The DNA and predicted protein sequences were different from previously known sequences. A hydrophobicity profile revealed the only handles to the putative role of the protein: It lacked a hydrophobic signal peptide but contained a single putative transmembrane domain after a hydrophilic 39 amino acid aminoterminal domain (Fig. 21.2). The only weak

Molecular Biology of Anhydrotic Ectodermal Dysplasia

413

Fig. 21.2. EDA molecule. Predicted protein sequence (A) and suggested transmembrane orientation of the product (B), including a summary of exon 1 mutations in the predicted EDA protein.

sequence homologies that were detected to any part of the EDA polypeptide involved the 75-residue hydrophilic carboxyterminal segment.3 It contained short runs of Gly-Xaa-Yaa repeats, the longest of which was four repeat units long. The overall structure, namely lack of a signal peptide and presence of a single transmembrane domain, suggested not only membrane localization but also orientation for the protein with the aminoterminus inside and carboxyterminus outside of the cell,55 and remains to be verified by antibodies directed toward different segments of the protein. The suggested type II orientation, presence of a single transmembrane domain, and collagenous repeats are reminiscent of a recently identified family of membrane-associated collagenous proteins.56,57 However, in contrast to these proteins, we have no evidence that the EDA protein would form homo- or heterotrimers such as collagenous proteins do, and presently EDA cannot be classified as belonging to this group. The functions of both true membranous collagens and the EDA protein are incompletely understood. Further analysis of patient DNA samples revealed that the putative gene was disrupted by both the AK and AnLy translocation breakpoints (the AnLy breakpoint mapped within the 200 kb intron) and that one or both exons were included in submicroscopic deletions in four male patients. A map of the X chromosomal region harboring the EDA gene is shown in Figure 21.3. Point mutations, including a de novo mutation, were detected in nine patients in exon 1.3 Mutations were discovered in only about 10% of patients studied, consistent with the length of exon 1 (638 bp) compared to the largest mRNA seen on Northern blots (6 kb). The location and type of known mutations in the predicted EDA protein are shown in Figure 21.2. Parallel studies on mouse DNA have further supported the concept that a homologous gene in mouse is responsible for the Ta phenotype. Nontranscribed as well as transcribed human DNA segments have revealed highly homologous counterparts in the mouse,54 and recently a comparative study has allowed us to identify the mouse Ta gene (A. K. Srivastava et al, in preparation). Its cloning should facilitate systematic functional and expression studies,

414

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 21.3. Map of the EDA gene region in the X chromosome. Yeast artificial chromosome (YAC) clones covering the region are shown above. A summary map indicates the positions of CpG islands (defined by restriction sites for the rare-cutter enzymes Not I [N], Eag I [E], BssH II [B] and Sac II [S]). The positions of exons 1 and 2 are indicated as double-headed arrows. Translocation breakpoints in female patients AK and AnLy and deletions in five male patients (ED series) are shown below.

and the gene’s identity could be directly confirmed by gene knock-out experiments (which should produce the familiar Ta phenotype) and gene replacement experiments (that should cure Ta mice).

EDA Expression and Function The cloning of EDA allowed the direct study of its expression pattern in the skin. Northern data and RT-PCR experiments suggested that EDA was expressed in many tissues, but in situ hybridization studies on adult skin scalp sections revealed a specific pattern.3 EDA mRNA was detected in the epidermis and epithelial cells in the upper hair follicle, sebaceous glands and, occasionally, eccrine sweat glands (Fig. 21.4). The lower hair follicle showed expression in the hair matrix and outer root sheath cells, whereas no signal was detected in the dermal papilla, dermis, or smooth muscle cells. These experiments revealed that EDA is an epithelial rather than a mesenchymal factor, even though our more recent results have also disclosed expression in some mesenchymal cell types.58 During the human fetal period, expression is seen in, besides epidermis, at least osteoblasts, neuroectodermal cells, thymus cells, and epithelial cells of the esophagus and kidney. In some cell types and tissues, such as keratinocytes, neurons, mammary gland, and prostate, expression continues through adult life.58 These observations correlate with the constant and occasional phenotypic features, such as facial and skull dysmorphism, thickening of esophageal and thymic epithelium, and occasional absence of nipples.20 The expression pattern also raises further questions for study, such as the role of the EDA gene product in adult tissues, especially skin and brain. Interestingly, the epithelia of the bronchi

Molecular Biology of Anhydrotic Ectodermal Dysplasia

415

Fig. 21.4. Expression of EDA in skin section. mRNA in situ hybridization results using a probe specific for EDA exons 1 and 2. (A) dark-field image of fetal skin from gestational week 20 shows signal in the epidermis and outer root sheaths in hair follicles (arrows). (B) an adult skin section shows signal in outer root sheath (arrowhead), sebaceous gland (arrow), and eccrine sweat gland tubules (curved arrows). (C) and (D) corresponding bright-field images, hematoxylin-eosin staining. Bar, 40 mm. See color insert.

and colon did not show specific signals. This result should be taken with caution, however, since it is possible that very low levels of expression were not detected by the in situ hybridization technique. The tissue types presently known to express EDA at some point during life are listed in Table 21.1. A single specimen of developing tooth at week 10 surprisingly showed absence of signal from the budding epithelium when the surrounding epithelium was positive by in situ hybridization, but then, again, a single specimen of adult tooth showed EDA mRNA in mesenchymal cells surrounding the tooth.58 At first, these observations seem puzzling because they do not conform to the pattern typical of most other morphogenetic factors. These results certainly need confirmation by additional and more targeted studies, soon to be possible using the mouse Ta gene probes and timed serial samples from developing wildtype and Ta mice. However, if confirmed, this pattern might correlate with the suggested multiple roles for the Ta gene observed in the early skin graft recombination experiments.30-32 Transient expression of EDA in cultured cells has allowed the verification of its suggested transmembrane localization.59 The 135 amino acid protein was expressed in monkey kidney COS-1 cells and biochemical fractionation and confocal imaging studies confirmed that the protein is translocated to the cell membrane (part of the protein was also found in endoplasmic reticulum). Future studies should address the effect of mutations on protein stability and translocation to the membrane. What, then, does such a small membrane protein do? With suggestive sequence homologies missing, many possibilities have to be kept open and explored by experimentation.

416

Molecular Basis of Epithelial Appendage Morphogenesis

Table 21.1. Human tissues known to express EDA mRNA and/or protein58 Cell Type

Expression in Fetal Period

Expression in Adult Tissue

Epidermis, keratinocytes Hair follicles Tooth bud, tooth Neuroectoderm, neurons Mammary epithelium Heart Thymus Osteoblast, bone Kidney epithelium Esophagus epithelium Prostate

yes yes (ectodermal cells) no yes n.d. yes yes yes yes yes n.d.

yes yes (ectodermal cells) yes (mesenchymal cells) yes yes no n.d. n.d. yes n.d. yes

n.d., no data.

A first hint came from further transient expression experiments using the human breast carcinoma cell line MCF-7.59 These cells normally grow flattened, attached to the culture dish surface. However, of those cells that were transfected with the EDA expression construct, more than 80% rounded up, lost the long actin filaments seen in nontransfected cells and lost contact with the dish surface except for a small area that still kept them attached 72 h after transfection. This effect was not seen in the COS-1 cells, perhaps suggesting that essential interacting factors are present in one but not in the other cell type. Any interacting proteins remain to be identified, but these observations offer a potential assay system for EDA function in cell culture. Mutagenized constructs can possibly be used to dissect functionally important domains of the EDA protein. Among potential roles for the EDA protein are that it might: 1. act as a locally secreted growth factor, analogous to the EGF family of growth factors; 2. act as a receptor or coreceptor, or possibly provide a phosphorylation target for receptors and thus participate in signal transduction; 3. participate in a cell’s contact with its surroundings by a role in intercellular or cellmatrix interactions; or 4. participate in membrane-cytoskeleton interactions. Of these possibilities, a role as a growth factor now seems slightly less likely, because no soluble product has been detected in the culture medium of cell lines transiently expressing high levels of the EDA protein (S. Ezer et al, work in progress). However, such a role is by no means excluded by current data. The functional role of the EDA protein may become first merged with the observations of reduced EGFR expression in EDA fibroblasts and change of cell shape in transiently overexpressing MCF-7 cells.34,59 Malignant cell lines that express abnormally high levels of EGFR and that upon stimulation by EGF round up and detach from a surface may provide useful reagents for further studies.60,61 Is the EDA protein involved in this phenomenon? Further knowledge of the function of the EDA gene will accrue from the study of its promoter elements and the regulation of its expression (G. Pengue et al, work in progress) and from the ongoing search for longer transcripts in human and study of possible alternative splicing. Even though we have mostly open questions rather than even partial answers, the cloning of the EDA gene has provided an entry to the elucida-

Molecular Biology of Anhydrotic Ectodermal Dysplasia

417

tion of mechanisms behind ectodermal dysplasias and the corresponding normal cellular functions.

Note Added in Proof After this review was submitted, two groups have characterized the mouse gene defective in the Ta mouse.62,63 The gene undergoes alternative splicing, and the longest deduced protein product is 391 amino acids.63 The protein contains a second, interrupted collagenous Gly-Xaa-Yaa domain and a cysteine-rich carboxyterminal domain. Novel homologous exons in human have been detected, extending the size of the human gene and protein.

Acknowledgments Our recent work on EDA has been the result of many individuals’ enthusiastic collaboration. I would like to especially thank the following friends and colleagues for discussions essential for writing this review and for continuing collaboration: Albert de la Chapelle, Sini Ezer, Johanna Pispa, Ulpu Saarialho-Kere, David Schlessinger, Anand K. Srivastava and Irma Thesleff. I also thank the anonymous reviewers for their helpful comments. Work in my laboratory is supported by the Sigrid Juselius Foundation, the Academy of Finland, the Finnish Pediatric Foundation (Ulla Hjelt Fund), and the Folkhälsan Institute of Genetics.

References 1. Darwin C. The Variation of Animals and Plants under Domestication Vol II. 2nd ed. London: John Murray, 1875:319. 2. McKusick VA. Mendelian Inheritance in Man. 11th ed. Baltimore: Johns Hopkins University Press, 1994. 3. Kere J, Srivastava AK, Montonen O et al. X-linked anhydrotic (hypohydrotic) ectodermal dysplasia is caused by mutation in a novel transmembrane protein. Nature Genet 1996; 13:409-416. 4. Thurnam J. Two cases in which the skin, hair and teeth were very imperfectly developed. Proc R M Chir Soc London 1848; 31:71-82. 5. MacKee GM, Andrews GC. Congenital ectodermal defect. Arch Dermat Syph 1924; 10:673. 6. Weech AA. Hereditary ectodermal dysplasia (congenital ectodermal defect). Am J Dis Child 1929; 37:766. 7. Smith J. Hereditary ectodermal dysplasia. Arch Dis Child 1929; 4:215. 8. Gordon WH, Jamieson RC. Hereditary ectodermal dysplasia of the anhydrotic type. Ann Int Med 1931; 5:358. 9. Thadani KI. The toothless men of Sind. J Hered 1934; 25:483-484. 10. Lord LW, Wolfe WD. Hereditary ectodermal dysplasia of the anhydrotic type (congenital ectodermal defect). Arch Dermat Syph 1938; 38:893. 11. de Silva PCC. Hereditary ectodermal dysplasia of the anhydrotic type. Quart J Med 1939; 8:97. 12. Sunderman FW. Persons lacking sweat glands. Hereditary ectodermal dysplasia of the anhydrotic type. Arch Intern Med 1941; 67:846-854. 13. Freire-Maia N, Pinheiro M. So-called “anhydrotic ectodermal dysplasia”. Int J Dermatol 1980; 19:455-456. 14. Freire-Maia N, Pinheiro M. Ectodermal dysplasias: A clinical and genetic study. New York: Alan R. Liss, 1984. 15. Freire-Maia N, Pinheiro M. Ectodermal dysplasias: A clinical classification and a causal review. Am J Med Genet 1994; 53:153-162. 16. Krafchik B. Ectodermal dysplasia. In: Alper JC, ed. Genetic Disorders of the Skin. St. Louis: Mosby Year Book, 1989:267-277. 17. Martin-Pascual A, De Unamuno P, Aparicio M et al. Anhydrotic (or hypohydrotic) ectodermal dysplasis. Dermatologica 1977; 154:235-243.

418

Molecular Basis of Epithelial Appendage Morphogenesis

18. Clarke A, Phillips DI, Brown R et al. Clinical aspects of X-linked hypohydrotic ectodermal dysplasia. Arch Dis Child 1987; 62:989-996. 19. Scaling skin in the neonate: A clue to the early diagnosis of X-linked hypohydrotic ectodermal dysplasia (Christ-Siemens-Touraine syndrome). The Executive and Scientific Advisory Boards of the National Foundation for Ectodermal Dysplasias, Mascoutah, Illinois. J Pediatr 1989; 114:600-602. 20. Reed WB, Lopez DA, Landing B. Clinical spectrum of anhydrotic ectodermal dysplasia. Arch Derm 1970; 102:134-143. 21. Kerr CB, Wells RS, Cooper KE. Gene effect in carriers of anhydrotic ectodermal dysplasia. J Med Genet 1966; 3:169-176. 22. Happle R, Frosch PJ. Manifestation of the lines of Blaschko in women heterozygous for Xlinked hypohydrotic ectodermal dysplasia. Clin Genet 1985; 27:468-471. 23. Happle R. Mosaicism in human skin. Arch Dermatol 1993; 129:1460-1470. 24. Hardy MH. The secret life of the hair follicle. Trends Genet 1992; 8:55-61. 25. Falconer DS. A totally sex-linked gene in the house mouse. Nature 1952; 169:664-665. 26. Grƒneberg H. Genes and genotypes affecting the teeth of the mouse. J Embryol Exp Morphol 1965; 14:137-159. 27. Blecher SR. Anhydrosis and absence of sweat glands in mice hemizygous for the Tabby gene: supportive evidence for the hypothesis of homology between Tabby and human anhydrotic (hypohydrotic) ectodermal dysplasia (Christ-Siemens-Touraine syndrome). J Invest Dermatol 1986; 87:720-722. 28. Grƒneberg H. The glandular aspects of the tabby syndrome in the mouse. J Embryol Exp Morphol 1971; 25:1-19. 29. Brockdorff N, Kay G, Cattanach BM et al. Molecular genetic analysis of the Ta25H deletion: evidence for additional deleted loci. Mammal Genome 1991; 1:152-157. 30. Sofaer JA. Differences between tabby and downless mouse epidermis and dermis in culture. Genet Res 1974; 23:219-225. 31. Mayer TC, Green MC. Epidermis is the site of action of tabby (Ta) in the mouse. Genetics 1978; 90:125-131. 32. Pennycuik PR, Raphael KA. The tabby locus (Ta) in the mouse: Its site of action in tail and body skin. Genet Res 1984; 42:51-63. 33. Blecher SR, Kapalanga J, Lalonde D. Induction of sweat glands by epidermal growth factor in murine X-linked anhydrotic ectodermal dysplasia. Nature 1990; 45:542-544. 34. Vargas GA, Fantino E, George-Nascimento C et al. Reduced epidermal growth factor receptor expression in hypohydrotic ectodermal dysplasia and tabby mice. J Clin Invest 1996; 97:2426-2432. 35. Miettinen P, Berger JE, Meneses J et al. Epithelial immaturity and multiorgan failure in mice lacking epidermal growth factor receptor. Nature 1995; 376:337-341. 36. Threadgill DW, Dlugosz AA, Hansen LA et al. Targeted disruption of mouse EGF receptor: effect of genetic background on mutant phenotype. Science 1995; 269:230-234. 37. Sibilia M, Wagner EF. Strain-dependent epithelial defects in mice lacking the EGF receptor. Science 1995; 269:234-238. 38. Cohen MM, Lin C-C, Sybert V et al. Two human X-autosome translocations identified by autoradiography and fluorescence. Am J Hum Genet 1972; 24:583-597. 39. MacDermot KD, Hulten M. Female with hypohydrotic ectodermal dysplasia and de novo (X;9) translocation. Clinical documentation of the AnLy cell line case. Hum Genet 1990; 84:577-579. 40. Turleau C, Niaudet P, Cabanis M-O et al. X-linked hypohydrotic ectodermal dysplasia and t(X;12) in a female. Clin Genet 1989; 35:462-466. 41. Limon J, Filipiuk J, Nedoszytko B et al. X-linked anhydrotic ectodermal dysplasia and de novo t(X;1) in a female. Hum Genet 1991; 87:338-340. 42. Shows TB, Brown JA. Human X-linked genes regionally mapped utilizing X-autosome translocations and somatic cell hybrids. Proc Natl Acad Sci USA 1975; 72:2125-2129. 43. MacDermot KD, Winter RM, Malcolm S. Gene localization of X-linked hypohydrotic ectodermal dysplasia (C-S-T syndrome). Hum Genet 1986; 74:172-173.

Molecular Biology of Anhydrotic Ectodermal Dysplasia

419

44. Kolvraa S, Kruse TA, Jensen PKA et al. Close linkage between X-linked ectodermal dysplasia and a cloned DNA sequence detecting a two allele restriction fragment length polymorphism in the region Xp11-q12. Hum Genet 1986; 74:284-287. 45. Clarke A, Sarfarazi M, Zonana J et al. X-linked hypohydrotic ectodermal dysplasia: DNA probe linkage analysis and gene localization. Hum Genet 1987; 75:378-380. 46. Hanauer A, Alembik Y, Arveiler B et al. Genetic mapping of anhydrotic ectodermal dysplasia: DXS159, a closely linked proximal marker. Hum Genet 1988; 80:177-180. 47. Zonana J, Clarke A, Sarfarazi M et al. X-linked hypohydrotic ectodermal dysplasia: Localization within the region Xq11-21.1 by linkage analysis and implications for carrier detection and prenatal diagnostics. Am J Hum Genet 1988; 42:75-85. 48. Zonana J, Jones M, Browne D et al. High-resolution mapping of the X-linked hypohydrotic ectodermal dysplasia (EDA) locus. Am J Hum Genet 1992; 51:1036-1046. 49. Jones AM, Malcolm S, Levinsky RJ et al. Physical mapping in the region of human Xq12-21.1 using pulsed field gel electrophoresis. Hum Genet 1993; 91:485-488. 50. Kere J, Grzeschik K-H, Limon J et al. Anhydrotic ectodermal dysplasia gene region cloned in yeast artificial chromosomes. Genomics 1993; 16:305-310. 51. Plougastel B, Couiliin P, Blanquet V et al. Mapping around the Xq13.1 breakpoints of two X/A translocations in hypohydrotic ectodermal dysplasia (EDA) patients. Genomics 1992; 14:523-525. 52. Zonana J, Gault J, Davies KJP et al. Detection of a molecular deletion at the DXS732 locus in a patient with X-linked hypohydrotic ectodermal dysplasia (EDA), with identification of a unique junctional fragment. Am J Hum Genet 1993; 52:78-84. 53. Thomas NST, Chelly J, Zonana J et al. Characterisation of molecular DNA rearrangements within the Xq12-q13.1 region, in three patients with X-linked hypohydrotic ectodermal dysplasia (EDA). Hum Mol Genet 1993; 10:1679-1685. 54. Srivastava AK, Montonen O, Saarialho-Kere U et al. Fine mapping of the EDA gene: A translocation breakpoint is associated with a CpG island that is transcribed. Am J Hum Genet 1996; 58:126-132. 55. Singer SJ. The structure and insertion of integral proteins in membranes. Annu Rev Cell Biol 1990; 6:247-296. 56. Pihlajaniemi T, Rehn M. Two new collagen subgroups: Membrane-associated collagens and types XV and XVII. Progr Nucl Acid Res Mol Biol 1995; 50:225-262. 57. Elomaa O, Kangas M, Sahlberg K et al. Cloning of a novel bacteria-binding receptor structurally related to scavenger receptors and expressed in a subset of macrophages. Cell 1995; 80:603-609. 58. Montonen O, Ezer S, Saarialho-Kere U et al.The gene defective in anhydrotic ectodermal dysplasia is expressed in the developing epithelium, neuroectoderm, thymus, and bone. J Histochem Cytochem 1998; 46:281-285. 59. Ezer S, Schlessinger D, Srivastava AK et al. Anhydrotic ectodermal dysplasia (EDA) protein expressed in MCF-7 cells associates with cell membrane and induces rounding. Hum Molec Genet 1997; 6:1581-1582. 60. Chinkers M, McKanna JA, Cohen S. Rapid rounding of human epidermoid carcinoma cells A-431 induced by epidermal growth factor. J Cell Biol 1981; 88:422-429. 61. Helseth E, Dalen A, Unsgaard G et al. Overexpression of the epidermal growth factor receptor gene in a human carcinoma cell line, derived from a brain metastasis. J NeuroOncol 1989; 7:81-88. 62. Ferguson BM, Brockdorff N, Formstone E et al. Cloning of Tabby, the murine homolog of the human EDA gene: Evidence for a membrane-associated protein with a short collagenous domain. Hum Molec Genet 1997; 6:1589-1594. 63. Srivastava AK, Pispa J, Hartung AP et al. Tabby phenotype is caused by mutation in the mouse homologue of the EDA gene, that reveals novel human exons and encodes a protein (ectodysplasin A) with collagenous domains. Proc Natl Acad Sci USA 1997; 34:1306-1307.

CHAPTER 22

Systematic Approach to Evaluation of Mouse Mutations with Cutaneous Appendage Defects John P. Sundberg, Xavier Montagutelli and Dawnalyn Boggess

Introduction

T

he inbred laboratory mouse has long been the species of choice for biomedical research including genetics based studies. In the past decade tremendous advances in molecular biology and gene manipulation have put the mouse into the forefront of species used in biomedical research. Large numbers of interesting mutations have been available for many years and are scattered around the world or maintained in repositories (see below). Transgenic and now targeted (so-called genetic “knockout”) mutations (collectively known as induced mutations) are being generated in large numbers, the most important of which are being collected and distributed from repositories. Access to these types of mice can be limited, but use depends upon the degree to which they have been characterized and compared to human diseases. This chapter will provide an approach to characterizing both existing and new mouse mutations and will summarize resources of information and repositories of mutant mice.

Systematic Evaluation of New Mouse Mutations Colony Establishment If a mouse is observed to be abnormal, it may be suffering from a common mouse disease that could be of an infectious nature or could be a common background disease characteristic of the inbred strain. Figure 22.1 provides a flow chart of how to proceed in evaluating this mouse. Laboratory animal veterinarians can often make the first decision as to whether this is a potential new mutation or a disease of importance to the colony or facility. Detailed descriptions of the common murine infectious diseases and strain specific diseases are published elsewhere.1-4 However, if only one mouse or a small group of mice within a litter are abnormal, these mice can be considered to be phenotypic deviants, or potential mutant mice. This discussion will focus on mutations that affect cutaneous appendages. The most common spontaneous mutations in mice are hair color, hair texture, and skin mutants in general, which can be due to hair or pilosebaceous defects (Table 22.1).5 If

Molecular Basis of Epithelial Appendage Morphogenesis, edited by Cheng-Ming Chuong. ©1998 R.G. Landes Company.

422

Molecular Basis of Epithelial Appendage Morphogenesis

Fig. 22.1. Example of a flow pattern to establish a colony and characterize the phenotype of a potential novel mouse mutation.

a mouse has an obvious defect, such as lack of hair or hair with a curly texture, this clinical feature (phenotype) can be followed in successive generations. Phenotypic deviants due to genetic mutations should be distinguished from those resulting from random events such as spontaneous diseases or accidental developmental defects. Evidence for genetic mutations include: 1. the occurrence of more than one mouse with the same abnormal phenotype within the same litter; 2. the recurrence of the same phenotype among mice from the same breeding pair but from different litters; 3. the occurrence of mice with the same phenotype in the progeny of different breeding pairs from the same pedigree; and/or 4. the presence in the same litter of both normal and affected mice. Once the genetic origin of the phenotype has been established, efforts should be undertaken to propagate it as a colony. The actual protocol will depend on the nature of the mutation (autosomal or sex-linked, recessive, semidominant, or dominant; see below) but

Evaluation of Mouse Mutations with Cutaneous Appendage Defects

423

Table 22.1. Examples of mouse mutations with skin appendage mutations5,42 Hair Fiber Abnormalities: Adrenocortical dysplasia (acd, Chr 8) Angora locus *(Fgf5go, Chr 5) Atrichosis (at, Chr 10) Balding locus (bal, Chr 18) Bareskin (Bsk, Chr 11) Caracul locus (Ca, Chr 15) Curly-whiskers (cw, Chr 9) Depilated (dep, Chr 4) Frizzy (fr, Chr 7) Fuzzy (fz, Chr 1) Hair interior defect (hid, not mapped) Hair patches (hpt, Chr 4) Harlequin (Hq, Chr X) Ichthyosis locus (ic, Chr 1) Lanceolate hair locus (lah, Chr 18) Lethal milk (lm, Chr 2) Matted (ma, Chr 3) Naked (N, Chr 15) Nude locus (nu, Chr 11) Ragged locus (Ra, Chr 2) Rex locus (Re, Chr 11) Satin (sa, Chr 13) Shaven (Sha, Chr 15) Silver (si, Chr 10) Soft coat (soc, Chr 3) Waved 1 (TγFαwa1, Chr 6) Waved 2 (Egfrwa2, Chr 11) Wellhaarig (we, Chr 2)

Eccrine Gland Abnormalities: Crinkled (cr, Chr 13) Downless locus (dl, Chr 10) Tabby locus (Ta, Chr X) Hair Cycle Abnormalities: Angora locus (Fgf5go, Chr 5) Hairless locus (hr, Chr 14) Hairy ears (Eh, Chr 15) Koala (Koa, Chr 15) Hair Follicle Adhesion Molecule Defects: Balding locus (bal, Chr 18) Lanceolate hair locus (lah, Chr 18) Nail Defects: Hairless locus (hr, Chr 14) Ichthyosis locus (ic, Chr 1) Sebaceous Gland Abnormalities: Asebia locus (ab, Chr 19) Crinkled (cr, Chr 13) Downless locus (dl, Chr 10) Hairless locus (hr, Chr 14) Harlequin ichthyosis (ichq, Chr 19) Tabby locus (Ta, Chr X)

* locus = more than one mutation (mulitple alleles) at this locus has been identified.

will always aim at maintaining the mutant allele together with the normal allele, so that at every step of the breeding process, control mice will be available that will be genetically as close to the mutants as possible. In most instances, inbreeding will be performed in order to increase the genetic homogeneity among individuals. In other cases, repeated crosses with an already established inbred strain (called recipient strain) will be made to “transfer” the mutation onto this particular background by the development of a congenic strain.6 If the mutation is autosomal dominant or semidominant, the matings should involve the crossing of an affected heterozygous mouse with a wild-type partner. In the case of inbreeding, the wild-type partner will be chosen from unaffected littermates. In the case of congenic strains, this will be a mouse from the recipient strain. In both cases, these crosses will yield affected and normal F1 mice in similar proportions (if the penetrance of the phenotype is full). The same protocol will be used for the next generations. If the mutation is autosomal recessive and the affected homozygous mice are fertile at least in one sex, inbreeding will be achieved by crossing a homozygous affected mouse with an obligate heterozygous partner. This cross will yield homozygous affected and heterozygous normal mice in similar proportions (if the penetrance of the phenotype is full). The same protocol will be used for the next generations. Creating a congenic strain will require

424

Molecular Basis of Epithelial Appendage Morphogenesis

a recurrent two-step process. First, a homozygous affected mouse will be mated with a partner of the recipient strain. They will produce only obligate heterozygous F1 mice. These will be intercrossed to produce 25% affected homozygous F2 mice which will be mated in turn with mice of the recipient strain, and so on. If the mutation is autosomal recessive and affected homozygous mice are sterile (or not viable) in both sexes, inbreeding will be achieved by crossing heterozygous breeders. Since these are phenotypically indistinguishable from wild-type mice, they will be identified by progeny testing as follows. Breeding pairs of phenotypically normal mice born from a breeding pair that yielded at least one affected mouse will be mated. Their progeny will be examined for the presence of affected mice. If no affected progeny are observed among the first 10 pups, it is very likely that at least one parent is not heterozygous. The breeding pair will then be discarded. On the contrary, in breeding pairs that yield at least one affected mouse, both parents are obligate heterozygous and are called a tested breeding pair. Their phenotypically normal progeny will be used to establish breeding pairs for successive generations. In the case of a congenic strain, a tested heterozygous breeder is mated with a partner of the recipient strain. Half of its F1 progeny will be heterozygous and will be identified by crossing with heterozygous tested breeders. Those F1 mice that yield at least one affected homozygous mutant will be mated with a partner of the recipient strain, and so on. If the mutation is sex-linked and affected hemizygous males or homozygous females are sterile, the colony will be maintained by crossing heterozygous females with wild-type males. In the case of recessive mutations, heterozygous females will be identified by progeny testing (heterozygous females are those which, when mated with wild-type males, yield affected males). For inbreeding, the males will be littermates of the heterozygous females. In the case of a congenic strain, they will be of the recipient strain. If affected hemizygous males and homozygous females are fertile, hemizygous males will be mated with heterozygous females at every generation in the case of inbreeding. In the case of a congenic strain, hemizygous males will be mated with females of the recipient strain. In the second generation, heterozygous females will be mated with males of the recipient strain. The third generation will be similar to the first one. When establishing a new inbred colony from unrelated individuals, it is considered that 20 generations of uninterrupted strict brother-to-sister matings are required in order to achieve a sufficient level of homozygosity (theoretically >99.9%). Ten crosses to the recipient strain should be performed for the establishment of a congenic strain. Mutations that are very difficult to maintain because affected mice are sterile or because pups die soon after birth can be maintained by ovarian grafts from mutants to oopherectomized severe combined immunodeficiency mutation (Prkdcscid/Prkdcscid) recipients who will accept grafts from nonhistocompatible strains.7 These females can then be mated with homozygous or heterozygous males.

Genetic Basis for the Phenotypic Deviant Assuming that a stable colony on the original inbred background has been established, the breeding data can be evaluated to look for trends that would indicate whether the new mutation is recessive or dominant, autosomal, or sex-linked. Dominant mutations give the same deviant phenotype in homozygous and heterozygous mice; recessive mutations give a deviant phenotype only in homozygous animals, whereas semidominant mutations show different deviant phenotypes in heterozygous and homozygous mice. All mutations with two classes of phenotypes in which affected mice are sterile are recessive in the mouse. X-linked mutations are either recessive (sometimes incompletely) or semidominant (in which case hemizygous males and homozygous females will show a similar phenotype,

Evaluation of Mouse Mutations with Cutaneous Appendage Defects

425

Fig. 22.2. Breeding patterns to illustrate recessive versus dominant autosomal mutations. Double boxes or circles indicate mutant mice.

whereas heterozygous females will show an intermediate phenotype). In X-linked mutations, the pattern of inheritance depends on the direction of the cross. If the mutation is semidominant, the cross of an hemizygous affected male with a wild-type female will yield heterozygous females and normal males, whereas the cross of a normal male with an homozygous female will yield affected hemizygous males and heterozygous females. If the mutation is recessive, the cross of an hemizygous affected male with a wild-type female will yield normal males, whereas the cross of a normal male with an homozygous female will yield affected hemizygous males and normal females. In the case of autosomal mutations, the mating of an affected mouse with a wild-type, unrelated partner will reveal the mode of inheritance of the mutation (Fig. 22.2). If this cross yields 100% affected F1 progeny, the mutation is dominant if the F1s resemble their parents, whereas it is semidominant if they show a different phenotype. If it yields 50% affected progeny, it is dominant if the crossing of F1s gives only one abnormal phenotype, whereas it is semidominant if two classes of deviant phenotypes are observed. Finally, the mutation is recessive in the case where all F1 progeny are normal. Crossing these F1 progeny will yield approximately 25% affected F2 progeny. These crosses should involve only laboratory strains to minimize the effect of background modifying genes on the phenotype. In the case of polygenic traits, the segregation pattern will often be more complex and will not fit with one of these simple models. The genetic analysis of these traits is beyond the focus of this overview.

Allelism Testing Once a stable colony has been established, another important step in characterizing a new mutant is to do allelism tests with other, previously characterized mouse mutations. Clinical and histological evaluation of the new mutant may reveal phenotypes that resemble other mutations. In this case, and only if the new mutation is recessive, it is possible to prove or to disregard the hypothesis that this mutation is a new allele (or remutation) at the same locus as one of the known mutations. The principle is that mice heterozygous for both mutations will show an abnormal phenotype if the two mutations affect the same gene, whereas they will appear normal if they affect different genes. Practically, either known heterozygous or homozygous mice of the new mutation are mated with either known heterozygous or homozygous mice of the known mutation. The observation that a fraction of the progeny show an abnormal phenotype close to that of either mutation demonstrates that the two mutations affect the same gene.

426

Molecular Basis of Epithelial Appendage Morphogenesis

When the map position of the new mutation is not known, allelism testing has to be done with every known mutation that resembles it. Alternatively, the genetic mapping of the mutation (see below) may suggest allelism with other mutations that are located in the same chromosomal region. In the case of semidominant or dominant mutations, proving allelism is much more difficult to achieve and will be based on either the occurrence of a new phenotype, or the frequency of the different classes of phenotypes, when crossing heterozygous mice.

Biological Characterization of a New Mutation Some believe that only a few mutant mice need to be studied, since inbred mice are essentially identical between individuals except for sex, that normal gross anatomy and histology are known makes controls unnecessary, and that a single gene mutation should be expressed in a repeatable manner between individuals. In a similar mind set, some believe that only a few of the organs, such as kidney, liver, and lungs, need to be examined to determine if there are systemic effects. These approaches can lead to erroneous and incomplete descriptions of mutant mice. Careful, systematic evaluation can be done to provide large amounts of information even when only small numbers of breeders are maintained. Mice, like other mammals, undergo major changes in their lives that may affect the phenotype of a mutation. These life changes can provide points to focus systemic investigations, while knowledge of aging changes of the organ system being studied will define a second set of observation points. Major life changes include birth (0-2 days), weaning (3-4 weeks), sexual maturity (6-8 weeks), sexual quiescence (6-8 months), and geriatric stages (2 years +) (Fig. 22.3). The age range provided reflects strain specific differences.8 The cost of maintaining mice to geriatric age is often prohibitive and many mice with genetic mutations never live that long, so this age group is rarely studied. Skin and its appendages undergo very dramatic changes during the first three weeks of life. The epidermis starts out relatively thick and becomes thinner by two weeks of age.9,10 The hair follicles are completing development during the first week of life, with the first hair fibers on the body emerging around five days of age.11 The hair cycles are synchronized and short during the first three weeks of life, so they can easily be studied. To evaluate all of these features, skin can be collected at two or three day intervals during the first three weeks of life.10 Numbers of mice can easily become quite large. If inbred mice are being studied and are maintained in a clean, environmentally controlled, pathogen free facility, as few as two mutants and two controls can be used of each sex (8 total per time point). Those collected for the major life changes should have complete sets of organs collected for study (see below) while those for skin and appendage evaluation can be limited to collection of selected skin. Use of the same individual for multiple biopsies can reduce the numbers of animals used but with skin mutants may cause a problem if the phenotype has a positive Köbner reaction (tape stripping to stress skin and exacerbate epidermal hyperproliferation), as is the case with some of the scaly skin mutations.12,13

Tissue Collection Injection of mice with bromodeoxyuridine or tritiated thymidine prior to euthanasia provides the opportunity to do kinetic studies later. Necropsy methods vary, but procedures for mice and other rodents have been detailed elsewhere.14 Mice can be euthanized by numerous methods depending upon institutional regulations. Carbon dioxide asphyxiation in a chamber is an American Veterinary Medical Association approved method for adults. Asphyxiation followed by decapitation is necessary for pups less than 12 days of age since they are rarely killed by the gas alone. Other methods are available and approved.7,15 Skin is

Evaluation of Mouse Mutations with Cutaneous Appendage Defects

427

Fig. 22.3. Major changes in the life of a mouse.

collected from the dorsal and ventral trunk, eyelids, ear, muzzle, tail, and footpads. Both vertical and horizontal sections are prepared for dorsal and ventral skin.16-18 Organs to collect include vertebrae, spinal cord, skeletal muscle, brown and white fat, femur, stifle joint, mammary gland, brain, liver, spleen, pancreas, intestines, stomach, kidney, adrenal, lung (inflated with fixative to dilate alveoli), urinary bladder, trachea, thyroid gland, esophagus, genital tract, heart, tongue and lymph nodes (Table 22.2). The anatomical locations and descriptions for the mouse are described in detail elsewhere.19 All tissues should be handled delicately to avoid crushing artefacts that will be obvious during microscopic evaluation. Tissue fixation is as critical as tissue collection. A large number of fixatives are available, each with their own advantages and disadvantages (Table 22.3). All organs should be routinely fixed by immersion. Skulls and long bones need to be decalcified following overnight fixation. These and other bony tissues are removed from fixative and placed in decalcifying solution, 50 parts decalcifying solution to 1 part tissue (Cal-Ex, decalcifying solution from Fisher Diagnostics, Fair Lawn, NJ). Tissues remain in the decalcifying solution overnight (up to 24 h) and are then trimmed, placed in marked cassettes and washed in running water for 3-4 h. After washing, the cassettes are placed in 70% ethanol and sent to the histology laboratory for processing. Soft tissues are fixed overnight, transferred into 70% ethanol, trimmed, processed, and embedded routinely in paraffin. Storage in ethanol is a critical step that stops crosslinking of amino groups in formalin fixed specimens, that, if allowed to continue, will mask many epitopes, making them useless for immunohistochemical studies. Blocks are sectioned at 5 µm and stained with hematoxylin and eosin for microscopic evaluation. Skulls are sectioned in three or more planes to evaluate the pituitary gland, eyes, ears, teeth, and other structures. Serial sections of skin can be stained with Masson’s trichrome (for scarring), Verhoeff-van Gieson (for elastic fibers), or periodic acid-Schiff (PAS; for fungi, glycogen, glycoproteins, and glycolipids), and Von Kossa (for mineralization) stains, among others.20,21 Skin can also be frozen for use in various biochemical, molecular, or immunofluorescence studies. Frozen sections lack detail but maintain epitopes for immunohistochemical studies or lipids that can be detected with oil red O, Sudan black, osmium, and other stains.20 We routinely take a 1.5 x 2.0 cm section of skin that is removed from the dorsal surface of the mouse at a standardized location, usually at the withers (tissue overlying the shoulders). This skin is trimmed to approximately 0.3 x 1.5 cm to form strips that run lengthwise from the head to the tail to properly orient hair follicles. The strips of skin are placed on a piece of aluminum foil, and a bead of OCT solution (Miles Inc., Elkhart, IN) is run along one edge. The foil is then dipped into liquid nitrogen to instantly freeze the samples. Once frozen,

428

Molecular Basis of Epithelial Appendage Morphogenesis

Table 22.2. Tissues to collect for detailed histopathologic overview of new mouse mutation. Normal, littermate controls should be collected at the same time for direct comparison. vertebrae spinal cord skeletal muscle femur stifle (knee) joint mammary gland liver spleen pancreas stomach kidney adrenal gland urinary bladder trachea thyroid gland genital tract heart tongue skull (includes pitutitary gland, ears, eyes, teeth, etc.)

brown and white fat brain intestines lung (inflate with fixative) esophagus lymph nodes

Table 22.3. Formulation of commonly used fixatives for preservation of mouse tissues. Tellyesniczky/Fekete’s acid alcohol formalin solution 100 ml of 70% ethanol 5 ml glacial acetic acid 10 ml formaldehyde solution (37% w/w) Modified Bouin’s Solution 85 ml of saturated aqueous picric acid 5 ml glacial acetic acid 10 ml formaldehyde solution Neutral Buffered 10% Formalin 100 ml formaldehyde solution 900 ml water 4 gm sodium phosphate monobasic 6.5 gm sodium phosphate dibasic

Carnoy’s Solution 60 ml absolute ethanol 30 ml chloroform 10 ml glacial acetic acid B-5 Fixative 6 gm mercuric chloride 1.25 gm sodium acetate (anhydrous) 98 ml hot distilled water Just prior to use add: 10 ml formaldehyde solution

samples are quickly removed from the foil and placed on edge in a clear plastic mold (HistoPrep disposable base molds, Fisher Scientific, Pittsburgh, PA) filled with OCT solution. The mold is then placed back into the liquid nitrogen and allowed to freeze. Frozen molds are wrapped in foil and stored at -80°C until needed. Both the molds and the foil are labeled with the accession number used for the animal being necropsied. In situ hybridization techniques can utilize tissues collected in fixatives routinely used for histopathology. However, use of these types of fixatives often is associated with high background signals. One approach that has yielded high quality results is the following. Strips of skin are collected in the same manner as for freezing described above. The skin is placed on nylon mesh, placed in 4% paraformaldehyde, and fixed overnight at 4°C. The following day the paraformaldehyde is replaced with 30% sucrose in phosphate buffered saline (pH 7.6) where it remains until the following day at 4°C. On day 3 tissues are removed, trimmed, and embedded in OCT solution as described above. These tissues can now be sectioned for in situ hybridization studies or shipped to collaborators on dry ice. Blood can be collected from the decapitation site of neonatal mice. Adult mice can have blood obtained from the retro-orbital venous plexus (so-called “eye-bleeding”), ventral coccygeal artery (so-called “tail-bleeding”), or cardiac puncture when the chest is opened.7 Sera can be collected from clotted, centrifuged samples for blood chemistry, electrolytes,

Evaluation of Mouse Mutations with Cutaneous Appendage Defects

429

and other analyses. Types of procedures and their interpretations in rodent samples are presented elsewhere.22 For molecular studies (gene mapping), spleen, liver, and kidneys are routinely removed at necropsy, snap frozen in liquid nitrogen in screw capped plastic vials (NUNC tubes, Nalgen Nunc International, Denmark), and stored at -80°C for subsequent DNA extraction.

Epidermal Kinetics Mutant and control (age and sex matched) mice can be injected intraperitoneally with 1 µCi/g body weight of tritiated thymidine (specific activity 20 µCi/mM; New England Nuclear, Boston, MA) or bromodeoxyuridine (50 µg/g body weight, Sigma Chemical Corp., St. Louis, MO) to label cells entering the “S” phase of the cell cycle. After one hour, mice are euthanized by CO2 asphyxiation, and complete necropsies performed. Skin is collected and fixed as described above, sectioned, covered with emulsion, incubated for 30 days, developed, and counterstained as previously described for tritiated thymidine.23,24 Bromodeoxyuridine incorporation is identified by immunohistochemistry. The number of positive nuclei (four or more silver grains over the nucleus for tritiated thymidine or dark brown nucleus for bromodeoxyuridine) are counted in the interfollicular epidermis per unit area or a defined number of nuclei. Positive cells are usually counted in a high dry (40 x) field. This can be applied to any anatomical structure in the skin or other organs studied.25

Cutaneous Morphometric Analyses The thickness of the interfollicular epidermis, various layers of the epidermis, dermis, hypodermal fat, and full skin can be measured with an ocular micrometer or much more simply and rapidly using a computer based image analyzer. Measurement of the hair follicle length can be used to demonstrate the hair cycle and shifts between mutant and control animals.10 Data can be put into spread sheets, analyzed, and graphically presented. Measurements of both dorsal and ventral truncal skin may reveal marked differences initially missed during histological review of slides.

Ultrastructural Evaluation Transmission electron microscopy can be a useful adjunct to standard histological studies. However, the relatively high cost of processing and viewing specimens warrants its judicious use. Skin is fixed for 18 h in 2.5% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.4, and postfixed for 18 h in aqueous 1% osmium or rhuthenium tetroxide.26,27 Tissues are stained en bloc with 2% uranyl acetate in 10% ethanol and further dehydrated in a graded ethanol series. Samples embedded in Spurrs-Mollenhauer resin are polymerized at 65°C for 48 h. Ultrathin sections are collected, stained with uranyl acetate and lead citrate, and examined in a transmission electron microscope.26 Scanning electron microscopy can be done on nails, skin punches, and plucked hairs. At the time of necropsy, 1.5-2.0 cm square samples of mouse skin can be removed from mutant and control mice, avoiding the subcutaneous fat layer. The samples are placed, connective tissue side down, on dry nylon mesh and immersed in cold 2.5% glutaraldehyde in 0.1 M cacodylate buffer. In addition, during necropsy the left front and rear feet can be amputated at the carpus/tarsus and prepared in a similar manner. After overnight fixation at 4°C, samples are washed twice with 0.1 m cacodylate buffer and postfixed in 0.5% osmium tetroxide in 0.1 m cacodylate buffer. Some of the skin samples can be first fractured in liquid nitrogen and then processed. Samples are subsequently dehydrated in a series of graded ethanols to 100%. After three changes in 100% ethanol the samples are critical point dried, attached to aluminum stubs with silver adhesive, and sputter coated with 15 nm of gold. Samples are examined in a scanning electron microscope operated at 10 kV. Plucked

430

Molecular Basis of Epithelial Appendage Morphogenesis

hairs from mice are directly mounted onto aluminum stubs with double stick tape. Hair samples are sputter coated with gold and examined as described above.26

Immunohistochemistry Serially sectioned skin can be screened with a variety of monoclonal and polyclonal antibodies. Careful search of the literature will define which antibodies work with mouse tissues and under what conditions. It is possible to set up standard methods and modify the dilution of the antibodies. Some antibodies require frozen sections while others work best in specific fixatives. Methods are described in detail elsewhere.23,28-30 Briefly, deparaffinized 6 µm serial sections are incubated in 3% H2O2 in methanol for 15 minutes to block endogenous peroxidase. A 30-minute incubation in 10% ovalbumin in phosphate buffered saline (pH 7.6) blocks nonspecific antibody adsorption. Slides are incubated overnight in primary antibody at 4°C, then washed 3 times in phosphate buffered saline. The reaction is detected using a modification of the avidin-biotin complex method (Vectastain ABC kit, Vector Laboratories) with diaminobenzidine (Sigma Chemical Corp.) as the chromagen. Mayer’s hematoxylin serves as the counterstain. Light green has been used successfully for screening and color photography, but is difficult to photograph with black and white film. Alkaline phosphatase kits with Histamark red (Kirkegaard & Perry, Gaithersburg, MD) can be used as well.

Skin Grafts Skin grafts can be useful to determine if the abnormal phenotype observed in the mutant mice is a primary or secondary event or, in the event that the mutation is a juvenile lethal, grafting can be used to determine if the cutaneous phenotype persists on recipients that live weeks or months beyond the normal life span of the mutant mouse.10,23,31 Briefly, donor mice are euthanized. Circular pieces of skin (epidermis and dermis), approximately 1 to 1.5 cm in diameter, are aseptically removed from the dorsal cervical and lumbar regions of the donor mice and placed in sterile phosphate buffered saline until the recipient mouse is ready to receive the graft. Recipient mice, either mice of the same inbred strain to avoid histocompatibility problems32 or immunodeficient mice such as nude (Hfh11nu/Hfh11nu)23 or severe combined immunodeficiency (Prkdcscid/Prkdcscid),10 are anesthetized with tribromoethanol (0.2 ml/10 g body weight; Aldrich Chemical Corp., Milwaukee, WI). A circular piece of skin (epidermis and dermis), approximately 1 to 1.5 cm in diameter, is aseptically removed from the left and right flanks of recipients. Donor skin is transplanted to these sites. Nexaban (Veterinary Products Laboratory, Phoenix, AZ) is applied to the apposing edges of the graft site to improve adherence. A piece of sterile, nonstick gauze pad (Telfa Pads, Kendall Co., Boston, MA) is cut to cover both grafts and is held in place with a strip of surgical tape (Micropore Surgical Tape, 3M Health Care, St. Paul, MN). Grafts can be further held in place with a self-adhesive crepe rubber bandage (3M Medical-Surgical Division, St. Paul, MN) cut to form a vest around the mouse. Each recipient receives one skin graft from a mutant and control donor mouse. In this manner the fate of each graft onto the same host can be followed. Each donor provides grafts for transplantation to two or more recipients.23

Mapping Mutant Gene Loci This is an important first step with spontaneous mutations to identify and investigate candidate genes that might be involved with the clinical phenotype described. This is not necessary for targeted mutations because the mutated gene is known and its localization has usually been established. The methodology is evolving rapidly and becoming easier and

Evaluation of Mouse Mutations with Cutaneous Appendage Defects

431

more efficient. Many approaches are available, only one of which will be briefly described here to provide the readers with a starting point. Intercrosses between two different inbred strains mixes the gene pool (Fig. 22.4). The more diverse the gene pools the greater the possibility for finding molecular polymorphisms between the inbred strains and therefore generating a high resolution map (Fig. 22.5, Table 22.4). The use of an intraspecific cross between inbred laboratory strains instead of using an inbred and a wild-derived inbred strain (an intersubspecific cross) has at least three advantages: 1. the genetic background might have a dramatic influence on the phenotype and, therefore, an intersubspecific cross could result in heterogeneous phenotypes that would make analysis more difficult; 2. there are currently nearly 6000 primers commercially available that detect simple sequence length polymorphisms, 40% of which are polymorphic for many combinations of standard inbred strains (depending on the method of analysis: horizontal agarose gel, nonradioactive acrylamide gel, or sequencing gel); and 3. intraspecific backcrosses can serve as the starting point for the establishment of congenic strains, thus saving time and animals, if results indicate such congenic strains would be of value. Using two inbred strains or an inbred strain crossed with an inbred wild strain, such as Mus musculus castaneus (CAST/Ei) in an intercross strategy, provides a first generation (F1) with no mutant offspring if it is a recessive trait. The second generation (F2), obtained by intercrossing the F1 mice, will have 25% offspring showing the mutation if it is due to an autosomal recessive gene defect (Figs. 22.2, 22.4). In those mice with the mutant phenotype, genes located immediately around the mutant locus will be homozygous for the allelic form present in the stock carrying the mutation. Conversely, the same mice will carry any combination of alleles at genes that are not genetically linked to the mutant locus. Therefore, using primer pairs for simple sequence repeats that can scan the mouse genome in polymerase chain reactions permits identification of these linked genes. Genomic DNA is isolated from spleen using standard procedures. Polymorphic microsatellites, previously published, are used in the polymerase chain reactions.33,34 Conditions for the primer pairs must be worked out for each parental strain of the cross being studied to generate clearly identifiable polymorphisms between the two parental strains. Analysis of F1 DNA provides a control (Fig. 22.5). Polymerase chain reaction products are all analyzed on 4% agarose gels (NuSieve™ 3:1 agarose, FMC Bioproducts, Rockland, ME) and stained with ethidum bromide to visualize the bands using a UV transilluminator. Linkage between the mutation and the molecular markers, genetic distances, and confidence intervals can be calculated using computer programs such as Gene-Link,35 MapManager36 or others. Markers can be ordered by minimizing double recombinants tested.

Resources and Repositories Informatics on Mouse Genetics Transgenic and targeted genetic technology provide powerful tools to induce mutations in laboratory animals, thereby creating new models for specific diseases and tools to dissect biological processes. Once valuable mutations are created, their use can be limited by access to information on their phenotypes as well as availability of the mice themselves. The Jackson Laboratory, a nonprofit research institution established in 1929, has developed the programs summarized here to meet these needs.37,38 The current Jackson Laboratory Informatics Program was formed in 1992. The Encyclopedia of the Mouse Genome software currently supports graphical displays of data derived

Molecular Basis of Epithelial Appendage Morphogenesis

432

Fig. 22.4. The harlequin ichthyosis mouse mutation was mapped by crossing BALB/cJ +/ichq females with CAST/Ei +/+ males to generate affected mice in the F2 generation.11

BALB/cJ

CAST/Ei

F1

Fig. 22.5. PCR analysis illustrates the DNA pattern for each parent (BALB/cJ or CAST/Ei) used to map the harlequin ichthyosis mutation.11 The offspring (F1) contain DNA from both parents.

Table 22.4. Simple sequence length polymorphisms (SSLPs) between BALB/cJ and other inbred mouse strains spanning 1.0-7.0 cM from the centromere on mouse Chromosome 19. Numbers of discernable polymorphic loci decrease as inbred strains increase in relatedness. A/J mice are derived from Bagg Albinos, the predecessor of BALB/cJ mice.8 Strain

SSLPs

Strain

SSLPs

Cast Spret Ob LP C57 NON

14 13 9 8 8 8

NON AKR DBA NOD C3H A/J

8 7 6 6 2 1

Evaluation of Mouse Mutations with Cutaneous Appendage Defects

433

from Chromosome Committee maps, MIT Genome Center SSLP maps, and maps generated directly from the Mouse Genome Database (MGD). MGD itself is a comprehensive resource of information about the mouse genome, including locus information, experimental genetic and physical mapping data, comparative mapping data for mouse, human, and 40 other mammals, probe, clone, and PCR primer information, genetic polymorphism data, phenotypic data including synoptic descriptions of genes (Mouse Locus Catalog) and inbred strain characteristics, and associated bibliographical references.39 MGD can be used for browsing and as a source of data to be analyzed and graphically displayed in the Encyclopedia of the Mouse Genome or other display tools. The MGD is available to the scientific community via the World Wide Web (WWW), where it is complemented by links to external databases such as On-line Mendelian Inheritance in Man (OMIM), Genome Database (GDB), and GenBank. MGD data are updated continuously, providing a unique and valuable resource for scientists interested in animal models and biomedical tools. The URL address for The Jackson Laboratory Bioinformatics server on the WWW is http:// www.informatics.jax.org. Mirror sites at the UK MRC Human Genome Mapping Project Resource Center in Cambridge, United Kingdom (http://mgd.hgmp.mrc.ac.uk), Institut Pasteur, Paris, France (http://www.pasteur.fr/Bio/MGD), and National Institute of Animal Industry/Japan Animal Genome Database in Ibaraki-ken, Japan (http://mgd.niai.affrc.go.jp) provide ready access to users in Europe and Asia, respectively.

Repositories of Mouse Mutations and Inbred, Congenic and Recombinant Inbred Strains The Jackson Laboratory Mutant Mouse Resource (MMR): 1. identifies and characterizes new spontaneous mouse mutations for biomedical research; 2. propagates new and established mouse mutations in useful stocks; 3. publishes information on these mutations; and 4. distributes mice carrying the mutations to scientists world wide. These include mice that are both found/created and characterized at The Jackson Laboratory as well as from laboratories of other scientists around the world. The MMR arose out of early mutant mouse characterization studies done by Drs. E.S. Russell and G.D. Snell in the 1930s and 1940s. This resource focuses on mutations that arise spontaneously, but also includes established mutations that were induced using radiation and mutagenic compounds such as ethylnitrosourea, as well as other classical approaches. Currently over 600 mouse mutations are available. Approximately 300 are maintained in small colonies, 300 as frozen embryos, and 30 in large production colonies.40 The Induced Mutant Resource (IMR) at The Jackson Laboratory was established in September 1992, in response to concerns from the scientific community about easy access to transgenic and targeted mutations. The IMR serves as a national clearing house for the collection and distribution of genetically engineered mice. The function of the IMR is to import, cryopreserve embryos, maintain, and distribute important transgenic, chemically induced, and targeted mutant strains of mice. Rederivation during importation rids mice of infectious diseases, and cryopreservation of embryos provides backup in case of accidental loss or intentional elimination of breeding colonies due to reduced demand. The IMR also undertakes genetic development of stocks, such as transferring mutant genes or transgenes onto defined genetic backgrounds (congenic strains) and combining transgenes and/or targeted mutations to create new mouse models for research. Currently over 400 induced mutants have been accepted into the program, many of which are currently being distributed world wide.41

434

Molecular Basis of Epithelial Appendage Morphogenesis

Acknowledgments This work was supported in part by grants from the National Institutes of Health (CA34196, RR08911, and AR43801) and the National Alopecia Areata Foundation. The authors thank J. Worcester and B.A. Sundberg for preparation of the computer generated graphics.

References 1. Lindsey RJ, Boorman GA, Collins MJ et al. Infectious Diseases of Mice and Rats. Washington, D.C.: National Academy Press, 1991. 2. Lindsey RJ, Boorman GA, Collins MJ et al. Companion Guide to Infectious Diseases of Mice and Rats. Washington, D.C.: National Academy Press, 1991. 3. Frith CH, Ward JM. Color Atlas of Neoplastic and Non-neoplastic Lesions in Aging Mice. Amsterdam: Elsevier, 1988. 4. Mohr U, Dungworth DL, Ward J et al. Pathology of the Aging Mouse. Washington, DC: ILSI Press, 1996. 5. Sundberg JP, King LE. Mouse mutations as animal models and biomedical tools for dermatological research. J Invest Dermatol 1996; 106:368-376. 6. Silver LM. Mouse Genetics: Concepts and Applications. New York, NY: Oxford University Press, 1995. 7. Cunliffe-Beamer TL. Biomethodology and surgical techniques. In: Foster HL, Small JD, Fox JG, eds. The Mouse in Biomedical Research. Vol. III. New York: Academic Press, 1983:401-437. 8. Green MC, Witham BA. Handbook on Genetically Standardized Jax Mice. 4th edition. Bar Harbor, ME: The Jackson Laboratory, 1991. 9. Sundberg JP, Boggess D, Sundberg BA et al. Epidermal dendritic cell populations in the flaky skin mutant mouse. Immunol Invest 1993; 22:389-401. 10. Sundberg JP, Rourk M, Boggess D et al. Angora mouse mutation: Altered hair cycle, follicular dystrophy, phenotypic maintenance of skin grafts, and changes in keratin expression. Vet Pathol 1997; 34:171-179. 11. Sundberg JP, Boggess D, Hogan ME et al. Harlequin ichthyosis (ichq): A juvenile lethal mouse mutation with ichthyosiform dermatitis. Am J Pathol 1997; 151:293-310. 12. Sundberg JP, Beamer WG, Shultz LD et al. Inherited mouse mutations as models of human adnexal, cornification, and papulosquamous dermatoses. J Invest Dermatol 1990; 95:62S-63S. 13. Nanney LB, Sundberg JP, King LE. Increased epidermal growth factor receptor in fsn/fsn mice. J Invest Dermatol 1996; 106:1169-1174. 14. Feldman DB, Seely JC. Necropsy Guide to Rodents and the Rabbit. Boca Raton, FL: CRC Press, 1988. 15. Close B, Banister K, Baumans V et al. Recommendations for euthanasia of experimental animals: Part 1. Laboratory Anim 1996; 30:293-316. 16. Headington JT. Transverse microscopic anatomy of the human scalp. A basis for a morphometric approach to disorders of the hair follicle. Arch Dermatol 1984; 120:449-456. 17. Whiting DA. The value of horizontal sections of scalp biopsies. J Cut Aging Cosmet Dermatol 1990; 1:165-173. 18. Whiting DA. Diagnostic and predictive value of horizontal sections of scalp biopsy specimens in male pattern androgenetic alopecia. J Am Acad Dermatol 1993; 28:755-763. 19. Popesko P, Rajtová V, Horák J. A colour atlas of the anatomy of small laboratory animals. Vol 2. Rat, Mouse, Golden Hamster. London: Wolfe Publishing Ltd., 1990. 20. Luna LG. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. New York: The Blakiston Division, McGraw-Hill Book Company, 1968. 21. Smith A, Bruton J. Color Atlas of Histological Staining Techniques. Chicago: Year Book Medical Publishers, Inc., 1977. 22. Loeb WF, Quimby FW. The Clinical Chemistry of Laboratory Animals. New York: Pergamon Press, 1989.

Evaluation of Mouse Mutations with Cutaneous Appendage Defects

435

23. Sundberg JP, Dunstan RW, Roop DR et al. Full thickness skin grafts from flaky skin mice to nude mice: Maintenance of the psoriasiform phenotype. J Invest Dermatol 1994; 102:781-788. 24. Smith RS, Hawes NL, Kuhlmann SD et al. Corn1: A mouse model for corneal surface disease and neovascularization. Invest Ophthalmol Vis Sci 1996; 37:397-404. 25. Dover R. Cell kinetics of keratinocytes. In: Leigh I, Lane B, Watt F, eds. The Keratinocyte Handbook. Cambridge: Cambridge Univ Press, 1993:203-234. 26. Morita K, Hogan ME, Nanney LB et al. Cutaneous ultrastructural features of the flaky skin (fsn/fsn) mouse mutation. J Dermatol 1995; 22:385-395. 27. Swartzendruber DC, Burnett IH, Wertz PW et al. Osmium tetroxide and ruthenium tetroxide are complementary reagents for the preparation of epidermal samples for transmission electron microscopy. J Invest Dermatol 1995; 104:417-420. 28. Sundberg JP, Burnstein T, Shultz LD et al. Identification of Pneumocystis carinii in immunodeficient mice. Lab Anim Sci 1989; 39:213-218. 29. Ueki H, Yaoita H. A Color Atlas of Dermatoimmunohistocytology. London: Wolfe Medical Publications Ltd., 1989. 30. Sundberg JP, Cordy WR, King LE. Alopecia areata in aging C3H/HeJ mice. J Invest Dermatol 1994; 102:847-856. 31. Hansen LA, Alexander N, Hogan ME et al. Genetically null mice reveal a central role for epidermal growth factor receptor in the maturation of the hair follicle and normal hair development. Am J Pathol (in press). 32. Gijbels MJJ, HogenEsch H, Bruijnzeel PLB et al. Maintenance of donor phenotype after full-thickness skin transplantation from mice with chronic proliferative dermatitis (cpdm/ cpdm) to C57BL/Ka and nude mice and vice versa. J Invest Dermatol 1995; 105:769-773. 33. Love JM, Knight AM, McAleer MA et al. Towards construction of a high resolution map of the mouse genome using PCR-analysed microsatelites. Nucleic Acids Res 1990; 18:4123-4130. 34. Dietrich W, Katz H, Lincoln SE et al. A genetic map of the mouse suitable for typing intraspecific crosses. Genetics 1992; 131:423-447. 35. Montagutelli X. Gene-link: A program in Pascal for backcross genetic linkage analysis. J Hered 1990; 81:490-491. 36. Manly KF. A Macintosh program for storage and analysis of experimental genetic mapping data. Mammal Genome 1993; 4:303-313. 37. Holstein J. The First Fifty Years at the Jackson Laboratory. Bar Harbor, ME: The Jackson Laboratory, 1979. 38. Sundberg JP. The Jackson Laboratory—A valuable medical research center. Comp Pathol Bull 1989; 21:1-4. 39. Nadeau JH, Grant PL, Mankala S et al. A rosetta stone of mammalian genetics. Nature 1995; 373:363-365. 40. Davisson MT. The Jackson Laboratory Mouse Mutant Resource. Lab Animal 1990; 19:23-29. 41. Sharp JJ, Davisson MT. The Jackson Laboratory Induced Mutant Resource. Lab Animal 1994; 23:32-40. 42. Sundberg JP. Handbook of Mouse Mutations with Skin and Hair Abnormalities: Animal Models and Biomedical Tools. Boca Raton, FL: CRC Press, 1994.

Index A Adhesion molecule 7, 9, 10, 16, 68, 77, 98, 102, 103, 123, 149, 150, 195, 231, 260, 265-267, 269, 270, 286, 307, 317, 320325, 330, 331, 333-335, 346, 357, 358, 388, 395, 397, 411, 423, 431 Afterfeather 62, 72 Anagen 63, 70, 76, 78, 82, 94, 95, 99, 100, 114, 122-125, 127, 128, 299, 301, 302, 305, 306, 309 Anhidrotic ectodermal dysplasia (EDA) 10, 15, 18, 20, 21, 23, 28, 30-32, 40, 41, 100, 172, 174, 410-419, 421, 424 Anterior foregut 203, 205-208, 210 Anterior-posterior axis 16, 69, 302 Antler 75, 111-114, 117, 118, 121, 124 Apical ectodermal ridge (AER) 24-27, 104, 136, 144, 168, 175, 274, 275, 282 Apoptosis 16, 22, 26, 70, 76, 82, 94, 96, 102, 119, 120, 169, 203, 211, 252, 271, 296, 297, 306, 308, 309, 316, 330, 419 Apteria 45, 46, 360

B Basement membrane (BM) 90-93, 157, 163, 164, 186, 221, 226, 231, 243-245, 258, 259, 284, 314, 332, 398 BMP-2 16, 25, 26, 92, 97, 98, 104, 164, 165, 168-171, 194, 249-251, 258, 259, 265, 269, 270, 272, 275, 278, 366, 367 BMP-4 16, 26, 27, 89, 91, 92, 97, 98, 101, 104, 164, 165, 167-170, 194, 195, 225, 226, 247, 249, 250, 256, 267-269, 271, 273277, 364, 366 Bone morphogenetic protein (BMP) 7, 16, 25-27, 89-92, 96-98, 101, 103, 104, 163, 164, 168-171, 225, 226, 247-250, 256, 266-269, 274-276, 302, 364, 366, 374 Bulge 11, 78, 80, 90, 93, 94, 102, 118, 119, 123, 293, 296

C Cadherin 7, 16, 88, 89, 93, 137, 187, 283, 285, 286, 293, 294, 296-298, 302-304, 342 Cadherin, E- (Ecad) 88, 89, 93, 137, 187, 285, 286, 293, 296-298, 302-304

Cadherin, P- (Pcad) 88, 89, 285, 286, 294, 296-298, 302-304 Catagen 70, 76, 78, 95, 114, 116-120, 122, 123, 287, 317 Catenin, beta- 285, 296, 302 Cell adhesion 7, 67, 68, 91, 217, 227, 228, 245, 252, 273, 277, 284, 286, 342, 348, 351, 352, 374, 400 Chorioallantoic membrane 5, 42, 47, 66, 401 Collagen 7, 21, 23, 67-69, 90, 91, 93, 137, 157, 158, 226-229, 251, 252, 255, 288, 289, 354, 362, 363, 413, 417 Craniosynostosis 21 Cutaneous appendage 3, 8, 45, 47-50, 52, 53, 395, 421. See also Skin appendage

D DCC (deletion in colorectal carcinoma) 195, 252 Dental lamina 161, 162, 166, 266, 269, 270 Dental papilla 26, 27, 157, 158, 163, 164, 169, 170, 269, 274 Dentin 157, 158, 266 Dermal condensation 47, 48, 52, 61, 62, 66-68, 90, 98, 244, 247, 251-253, 255258, 297-299, 359, 361-363 Dermal papilla 10, 29, 57-59, 63, 70, 76, 79, 80, 83, 88-90, 92-96, 98, 100, 102-104, 140, 256, 291, 293, 294, 303, 316, 322, 323, 328, 414 Dermis 40, 45-50, 52, 53, 57, 62, 64, 66, 67, 69, 70, 78-80, 82, 83, 88, 91-93, 96, 97, 100, 113, 117, 119, 136, 137, 140-142, 145, 146, 247, 251, 252, 254, 256, 293, 294, 298, 299, 322, 359, 361-363, 411, 414, 429, 430 Dlx 7, 21, 23, 162, 273 Dorso-ventral axis 23-25, 45, 53, 145, 203, 208 Dpp 187, 189, 194, 225 Drosophila 5, 7, 23, 28, 50, 98, 102, 112, 116, 121, 161, 164, 166, 170, 182, 186-190, 193-195, 204-208, 217, 220, 222, 223, 225, 226, 249, 250, 268, 271, 273, 275, 346, 348, 350, 353, 354, 360, 361, 374, 404

442

Molecular Basis of Epithelial Appendage Morphogenesis

E EGFR 89-91, 93, 99, 222, 223, 231, 270, 411, 416 Enamel 27, 28, 95, 157, 158, 163, 164, 166170, 172, 173, 266, 269-271, 274, 275 Epidermal growth factor (EGF) 23, 27-30, 90, 91, 93, 99, 100, 123, 166, 172, 218, 222, 223, 229, 251, 266, 270, 285, 328, 411, 416 Epithelial appendage 3-12, 15, 30, 58, 62-64, 72, 157, 158, 162, 163, 166, 169, 172, 173, 215-217, 220, 224, 243, 332, 366, 371, 395, 396 Epithelial placode 79, 83, 88-90, 92-96, 98-101, 104, 245, 247, 249, 250, 255, 350, 371 Epithelial-mesenchymal interaction 4, 8, 11, 20-24, 26-28, 30, 72, 92, 100, 102, 103, 111, 162-164, 166, 171-173, 181, 207, 209, 217, 218, 228, 243, 244, 247, 251, 252, 256, 265-268, 271-277, 298, 378, 410 Epithelial-mesenchymal recombination 67, 247, 362, 383, 404 Epithelioblast 10-12 Epithelioid 343, 344, 346, 352, 354 Epithelium 5, 17, 20, 25-27, 29, 58, 60, 64, 66-69, 71, 72, 75, 78, 83, 88, 89, 92-94, 97, 98, 100-104, 119, 123, 131, 132, 136, 157, 158, 160-174, 181, 184-187, 189, 192, 193, 196, 206, 207, 209, 215-219, 221-224, 226, 230, 243-245, 247, 249255, 258, 266, 267, 269-276, 288, 291, 297, 299, 300, 302, 304, 362, 397, 399, 404, 414-416 Esophagus 184, 187, 192-194, 203, 206-208, 414, 416, 427, 428 Explant culture 249, 297, 298, 364, 399, 400, 404 Extracellular matrix 16, 22, 23, 27, 30, 67, 77, 90-93, 96, 119, 136, 158, 163, 164, 186, 217-219, 225-227, 229, 232, 251, 253, 266, 273, 284, 287, 353, 355, 362-364, 374

F Feather 3, 5, 7-10, 16, 41, 42, 45-50, 52, 53, 57-72, 75, 77, 78, 83, 93, 95, 97, 98, 100, 103, 114, 124, 140, 141, 157, 169, 243258, 283, 296, 298, 299, 301, 304, 315317, 331, 332, 350, 355, 359-364, 366, 367, 371, 383, 386, 387, 395, 399-402, 405

Feather, barb 49, 57, 58, 61, 62, 64, 71, 72, 140, 141, 331, 387, 402 Feather bud 5, 7, 48, 58, 60-63, 66-71, 95, 98, 100, 140, 243-258, 301, 331, 359, 361-364, 366, 399, 400 Feather follicle 63, 64, 70, 71, 298, 301, 315, 316, 331, 332 Feather keratin 71, 316, 317, 331 Feather primordia 52, 60-62, 64, 67-70, 244, 249, 251, 254-256, 360, 362, 364, 366 Feather tract 46, 58, 61, 62, 66, 255, 366 FGF receptor (FGFR) 17, 22, 70, 89, 90, 92, 100, 220-223, 231, 247, 250, 300, 302 FGF-1 89, 92, 100, 245, 274 FGF-2 24, 26, 47-49, 89, 92, 100, 170, 245 FGF-4 16, 24-26, 92, 100, 103, 164, 166-169, 172, 194, 245, 247, 249, 250, 269, 270, 274, 275, 364 FGF-5 70, 120 Fibroblast growth factor (FGF) 7, 11, 16, 22-26, 46-49, 68-70, 89-92, 100, 103, 104, 118, 120, 122, 137, 164-170, 172, 194, 218, 220, 222, 223, 226, 245, 247, 249, 250, 256, 257, 265-267, 269, 270, 273-275, 277, 302, 364, 366, 374, 404, 423 Fibronectin 7, 68, 69, 91, 137, 163, 225, 251, 252, 287-289, 363 Forkhead (fkh) 189, 195, 206, 220, 222

G Gene supernetwork 371, 372, 379, 381-387 Gene therapy 10, 232 Gene totinetwork 385 Gland 3, 5, 8, 17, 18, 20, 22, 23, 28-30, 40, 41, 45, 49, 75, 78, 80, 82, 96, 99, 100, 112, 114, 119, 121, 122, 166, 170, 172, 183186, 203, 207-209, 211, 216, 220, 228, 266, 272, 286, 290, 293, 296, 302, 315, 322-324, 346, 347, 359, 371, 410-412, 414, 415, 423, 427-429 Growth factors 16, 23, 53, 92, 111, 119, 122124, 137, 145, 147, 164, 221-223, 229, 244, 245, 248, 250, 252, 253, 256, 265-269, 271, 273-276, 364, 400, 416 Gut 3, 7, 23, 111, 112, 142, 143, 181-184, 186-189, 192-196, 203-210, 215-217, 222, 249, 268, 271, 421

H Hair cortex 315, 318, 322, 330 Hair cuticle 315, 320, 324-326 Hair cycle 70, 78, 82, 94, 111, 116-118, 121123, 142, 317, 402, 423, 426, 429

Index Hair follicle 9, 12, 17, 20, 23, 27, 29, 39-42, 45, 52, 70, 71, 75-80, 82-84, 88-104, 111, 114, 115, 117, 121-123, 138, 140, 145, 170, 266, 271, 272, 283, 285, 286, 288, 290, 291, 293-295, 298, 301, 303, 315317, 322-324, 327, 328, 330-332, 363, 411, 414-416, 423, 426, 427, 429 Hair germ 78-90, 92-94, 99, 101, 103, 104, 117, 286, 298 Hair peg 78-80, 89, 93, 94, 286, 298, 299 Hair pelage 48, 49, 53 Heterochrony 72, 341 Homeobox 7, 16, 20, 23, 24, 26, 50, 53, 89, 95, 162, 172, 187, 189, 205, 245, 250, 265, 267, 271, 275, 291, 292, 301 Homeodomain 20, 21, 26, 172, 189, 205-208, 267, 272, 301 Hox 7, 16, 23, 50-53, 89, 95, 98, 104, 161, 189, 190, 192-194, 205, 245, 269, 287, 291, 292, 301, 302, 366, 374, 377 Hypodontia 15, 18, 19, 22, 172, 409

I Immunoglobulin superfamily 283, 285 Integrin 7, 16, 23, 68, 91, 99, 219, 251, 252, 283, 285, 287-290, 296, 297, 300, 301, 304, 363 Intestine 3, 184-187, 192, 427, 428

K Keratin 19-21, 30, 41, 42, 48, 71, 80, 82, 91, 93, 97, 99, 101, 122, 315-317, 319, 320, 322-325, 327-332 Keratinocyte 9, 11, 83, 90, 96, 99, 100, 137, 140, 141, 270, 285, 288, 289, 293-295, 303, 315, 317, 324, 328, 330, 332, 414, 416

L Laminin 137, 227, 252, 288, 289, 346 Lef1 20, 23, 27, 28-30, 89, 92, 95, 97, 101, 104, 170, 171, 265, 267, 271, 272, 274, 276, 277, 285, 302, 327-329 Limb bud 7, 24-27, 29, 30, 53, 66, 68, 75, 98, 100, 113, 182, 189, 192, 194, 217, 245, 250, 268, 271, 275, 341, 371, 385 Lmx 7 Lung 3, 8, 20, 23, 30, 169, 181, 182, 194, 203, 205-211, 215-232, 243, 245, 266, 346, 426-428

443

M Melanoblast 17, 131-134, 136, 137, 140-147 Melanocyte 9, 17, 64, 76, 117, 131-134, 136-138, 140-147, 287, 291 Mesenchyme 3, 5-7, 17, 18, 20, 24-27, 29, 30, 51, 53, 62, 64, 66, 67, 69, 72, 75, 79, 83, 88-93, 95-98, 101-104, 113, 123, 131, 132, 137, 157, 158, 160-164, 166, 168173, 185, 186, 207-209, 215, 217-219, 222, 223, 226, 232, 243-245, 247, 249-255, 258, 266, 267, 269, 270-277, 297, 298, 300, 361, 362, 397, 399 Mesoderm 16, 17, 23-26, 46, 48, 49, 78, 82, 104, 133, 160, 162, 181-187, 189, 192195, 205, 243, 245, 249, 267-270, 275, 289, 341 Metastasis 12 Morphogen 164, 217, 268, 291, 292, 295, 299, 300, 364 Morphogenesis 5-7, 9, 10, 12, 16, 18, 23, 24, 26, 29, 30, 46, 47, 49-53, 58, 61, 62, 64, 65, 68, 71, 75-78, 83, 88, 93, 94, 96, 97, 102-104, 115, 157-159, 161-164, 166, 169-173, 181, 184, 187-189, 192-194, 203-211, 215-232, 243-245, 249-253, 257, 258, 265-270, 272-274, 276, 283289, 291, 293-305, 331, 341, 342, 344, 346-348, 352-355, 362, 363, 378, 381383, 395, 396, 401, 404, 406, 410 Msx-1 16, 17, 20-23, 26, 27, 30, 89, 95, 97, 170-172, 245, 249, 250, 265, 267, 271-277 Msx-2 16, 17, 21, 23, 26, 27, 89, 95, 97, 171, 172, 245, 249, 250, 265, 271-275

N NCAM 7, 66, 68, 69, 71, 91, 93, 137, 245, 252, 253, 255-258, 285-287, 289, 290, 295, 296, 298-304, 363 Neural crest 5, 9, 17, 20, 26, 30, 46, 66, 82, 101, 102, 131-134, 136, 137, 140, 142145, 147, 157, 158, 160-162, 173, 266, 269, 271, 272, 277 Neurotrophin receptor 91, 166 Notch 69, 90, 102, 103, 166, 245, 255, 323, 361

O Odontoblast 157, 158, 163, 164, 166, 169, 266 Oncogene 139, 255, 346, 403 Otx 50

444

Molecular Basis of Epithelial Appendage Morphogenesis

P Paired box (Pax) 170, 171, 196, 211, 265, 267, 271, 273, 287, 291, 301, 302 Paralogue 192 Pattern formation 24, 28, 30, 58, 68, 77, 78, 95, 145, 189, 203-205, 208, 209, 244, 249, 270, 284, 285, 287, 291, 297, 298, 301303, 305, 345, 347, 348, 350, 352, 353, 359-362, 364, 366 Patterning 5, 9, 23-25, 27, 28, 30, 45, 47, 50, 67, 68, 71, 95, 137, 138, 161, 162, 168, 169, 173, 186, 187, 189, 192-194, 206, 226, 247, 249-251, 359, 360, 362-364, 367 Pigment 131-138, 140-147, 348 Placode 5, 16, 29, 30, 45, 47, 49, 60, 62, 64, 66, 67, 69, 79, 83, 88-90, 92-104, 220, 245, 247, 249, 250-253, 255, 271, 290, 291, 293-296, 298, 300, 302, 350, 353, 359, 361, 371, 385 Polarity 23, 24, 98, 193, 226, 284, 285, 291, 292, 346, 361, 378 Proteoglycan 23, 68, 90, 91, 120, 122, 123, 137, 164, 226, 227, 229, 251, 253, 273, 363 Proximal-distal axis 45, 70 Pteryla 45, 46, 58, 359-361, 364

R RCAS 7, 247, 253, 403, 404 Recombination 5, 6, 46, 48, 52, 67, 69, 72, 83, 88, 136, 143, 160, 162, 163, 170, 182, 206, 207, 217, 243, 247, 250, 253, 266, 269, 272-274, 362, 374, 383, 399, 404, 411, 415 Retinoic acid (RA) 5, 7, 49, 53, 95, 96, 102, 161, 270, 330, 400, 401 Retinoic acid receptor (RAR) 49, 90 Retroviral vector 194, 247, 297, 401, 403

S Secondary enamel knot 168, 169 Segmentation 184, 326, 348, 350-352 Signaling 7, 9, 11, 16, 23-30, 46, 53, 66, 68, 71, 77, 78, 88-90, 92, 94-96, 98-100, 102-104, 112, 147, 162, 164, 166-173, 181, 182, 189, 193, 194, 196, 204, 205, 217-219, 222-226, 230-232, 244, 251253, 255, 256, 265, 267, 269-277, 287, 297-299, 363, 364, 366 Skin appendage 5, 7, 62, 66, 68, 70, 71, 83, 95,

96, 98, 100, 104, 244, 247, 253, 298, 301, 364, 372, 385, 390, 391, 403, 406, 410, 415. See also Cutaneous appendage Sonic hedgehog (Shh) 7, 11, 16, 24-28, 60, 68, 69, 71, 92, 98, 100, 104, 166-169, 172, 193, 194, 226, 245, 247, 249, 250, 252, 253, 255-258, 266, 267, 270, 271, 274, 364, 366, 404 Stem cell 10, 11, 30, 80, 91, 94, 119, 138, 140, 161, 230, 285, 291, 293, 296 Stem cell factor (SCF) 139-142, 145, 146

T Tabby 27, 28, 99, 100, 172, 173, 409, 411, 423 Teeth 3, 5, 8, 10, 15, 17, 18, 20-23, 27, 28, 75, 99, 157-167, 169-173, 243, 245, 271, 272, 302, 371, 383, 409-411, 427, 428 Telogen 70, 76, 78, 114, 116, 118, 119, 121, 122, 299, 301 Tenascin 23, 68, 69, 90-93, 95, 137, 164, 184, 251, 253, 256, 258, 273, 274, 277, 288, 289, 300, 363 TGF-α 91, 99, 100, 166, 222, 270, 328 TGF-β 23, 26, 68, 83, 84, 90-92, 96, 97, 99, 104, 123, 137, 164, 187, 221, 222, 224226, 230-232, 249, 252, 253, 255-258, 266, 267, 269, 270, 274, 275, 287, 302, 328 Thyroid 71, 118, 122, 181, 192, 203, 205-211, 221, 328, 427, 428 Tooth development 7, 21, 27, 29, 32, 95, 101, 157-165, 170-172, 265-267, 269-277, 289 Topobiology 9, 283, 284, 288, 291, 295, 296, 298, 302-305 Trachea 203, 206-208, 217-223, 427, 428 Transcription 7, 16, 17, 20, 23, 26, 27, 29, 30, 46, 53, 77, 89, 98, 101, 102, 104, 116, 147, 162, 169-171, 187, 189, 195, 203-211, 217-222, 225, 226, 231, 244, 247, 249, 250, 255, 265-268, 271-274, 283, 285, 301, 302, 315, 327-331, 350, 352, 377, 378, 381, 403

W Wnt 7, 23-26, 53, 69, 170, 226, 245, 250, 267, 374 Wool 40-42, 77, 114, 115, 121, 122, 319-321, 324-327, 362, 363

Z Zone of polarizing activity (ZPA) 16, 24-26, 28, 168, 169, 194, 274, 275