Systematic Evaluation of the Mouse Eye: Anatomy, Pathology, and Biomethods (Research Methods For Mutant Mice)

Systematic E va l u a t i o n o f t h e MOUSE EYE A n a t o m y, P a t h o l o g y, and Biomethods Research Methods ...

Author: Richard S. Smith | Simon W. M. John | Patsy M. Nishina | John P. Sundberg

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Systematic E va l u a t i o n o f t h e

MOUSE EYE A n a t o m y, P a t h o l o g y, and Biomethods

Research Methods for Mutant Mice Series Series Editor

John P. Sundberg Systematic Approach to Evaluation of Mouse Mutations John P. Sundberg and Dawnalyn Boggess

Systematic E va l u a t i o n o f t h e

MOUSE EYE A n a t o m y, P a t h o l o g y, and Biomethods

editor-in-chief

Richard S. Smith co-editors

Simon W. M. John • Patsy M. Nishina John P. Sundberg Graphics and Artwork by Jennifer L. Smith

CRC PR E S S Boca Raton London New York Washington, D.C.

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Cover Art Front Cover. Top. Anterior chamber angle, trabecular meshwork, and Schlemm’s canal of adult mouse. Bottom. Choroidal neovascularization extends into the subretinal space and retina in a Bst mouse. Back Cover. Diagram showing dissection of the anterior segment for fixation and embedding for transmission electron microscopy. Senior Editor: John Sulzycki Project Editor: Christine Andreasen Production Manager: Carol Whitehead Marketing Manager: Nadja English Cover Designer: Dawn Boyd

Library of Congress Cataloging-in-Publication Data Systematic evaluation of the mouse eye: anatomy, pathology, and biomethods / editor-in-chief, Richard S. Smith; co-editors, Simon W. M. John, Patsy M. Nishina, John P. Sundberg. p. cm. -- (Research methods for mutant mice series) Includes bibliographical references. ISBN 0-8493-0864-X (alk. paper) 1. Mice--Anatomy. 2. Eye. I. Smith, Richard S. (Richard Stanton), 1934- II. Series. QL737.R6 S96 2001 573.8′81935--dc21

2001052417

This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming, and recording, or by any information storage or retrieval system, without prior permission in writing from the publisher. All rights reserved. Authorization to photocopy items for internal or personal use, or the personal or internal use of specific clients, may be granted by CRC Press LLC, provided that $1.50 per page photocopied is paid directly to Copyright Clearance Center, 222 Rosewood Drive, Danvers, MA 01923 USA. The fee code for users of the Transactional Reporting Service is ISBN 0-8493-0864-X/02/$0.00+$1.50. The fee is subject to change without notice. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. The consent of CRC Press LLC does not extend to copying for general distribution, for promotion, for creating new works, or for resale. Specific permission must be obtained in writing from CRC Press LLC for such copying. Direct all inquiries to CRC Press LLC, 2000 N.W. Corporate Blvd., Boca Raton, Florida 33431. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation, without intent to infringe.

Visit the CRC Press Web site at www.crcpress.com © 2002 by CRC Press LLC No claim to original U.S. Government works International Standard Book Number 0-8493-0864-X Library of Congress Card Number 2001052417 Printed in the United States of America 2 3 4 5 6 7 8 9 0 Printed on acid-free paper

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Preface With completion of the first phase of the Human Genome Project, it is evident that scientists are confronted with mountains of information. The availability of all human genes and their locations is an exciting prospect. However, the nature of the proteins coded for by these genes and their molecular mechanisms of action are often unknown. Efforts to take advantage of this new fund of knowledge are complicated by several factors: (1) every human has a unique set of genes that make the search for cause and effect a challenging endeavor; (2) infinite variations in environment influence gene expression in humans and such factors are difficult to control; (3) gene expression in humans is difficult to study because many tissues and organs are not easily or safely accessible for biopsy; (4) tissue from diseased human organs is sometimes obtained only after years of impact from medicine, surgery, and the disease itself; and (5) many diseases are complex traits and are the result of the interaction of many genes, some of which exert modest effects; such genes are difficult to detect or map. These problems are best addressed by mouse genetics. It is well known that 90 to 95% of the mouse and human genomes is shared. This fact makes it likely that a given gene in a mouse will be found in a homologous chromosomal segment in humans and will exert its effects through similar mechanisms. The first draft of the mouse genome is already available. The list of advantages in using mice for genetic research is long: (1) Inbred strains of mice are genetically identical except for the mutation being studied; (2) evolution of disease is more rapid in mice because of their 2- to 3-year life span; (3) diet and environment can be precisely controlled; (4) tissue is available for study early in the disease process; (5) there are many well-characterized spontaneous mutant mice available; and (6) genes can be manipulated by targeted mutagenesis. Although numerous sophisticated techniques are available in genetics, the study of disease remains firmly linked with the disciplines of anatomy and pathology. In some tissues, structural details are relatively simple and easily interpreted. The eye, however, is an asymmetrical organ of complex structure with important regional anatomic variations. If orientation of the eye during sectioning is not carefully controlled, it is easy to make erroneous interpretations. This problem is complicated by the size of the mouse eye and the fact that the eyes of all animals are challenging tissues to fix and section without artifacts. These difficulties can be overcome by use of the techniques discussed in this book. Our primary goals are to describe the normal anatomy, development, general pathology, and methodology for evaluation of the mouse eye and adnexae. Although this information is available in the scientific literature, it is widely scattered. Efforts have been made to provide the reader with extensive bibliographies at the close of each chapter as resources for additional information. A separate CD that contains all figures in color is included with the book, because color versions of the figures offer details not obvious in black-and-white pictures. Where needed, electron microscopic pictures illustrate features not visible with light microscopy, although this book is not intended to be a comprehensive guide to ocular ultrastructure. The middle sections of the book review issues in mutant evaluation and the ocular histopathological features of diseases that occur in response to genetic and environmental stimuli. Space does not permit a complete review of the histopathological features of every genetically engineered mouse that has been reported; the material presented is intended to cover the general morphological changes in mice and to provide a basis for interpreting new mouse mutations. A constant effort has been made to relate human ocular disease to mouse models of disease. Chapters on mapping of complex traits and on ENU mutagenesis are included as an introduction to these important techniques and their relationship to analysis of morphology.

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The final portion of the book is devoted to procedures for tissue preparation and interpretation as well as to methodology useful for examination and evaluation of the mouse eye. It is difficult to think of doing genetic research without the huge reservoir of information available online, so the final chapter describes Internet resources that provide supplementary information. Richard S. Smith

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Acknowledgments Although the editors and authors have provided many of the illustrations in this work, we could not have produced this book without the willingness of many scientists to share their material with us. To those who have shared slides, photographs, or copies of their previously published materials, we express our deepest thanks. Their contributions are acknowledged throughout the book. We are, of course, grateful to the publishers of journals and books who have agreed to let us use previously published materials. Nearly all of the original material published here is a product of numerous research efforts both at The Jackson Laboratory and at the institutions of the contributing authors. This project would not have been possible without the generous financial support of the Howard Hughes Medical Institute (Simon W. M. John is an assistant investigator and Richard S. Smith is a research specialist of HHMI at The Jackson Laboratory); the National Cancer Institute (for Cancer Center Core Grant P3034196); the National Eye Institute (multiple grants), and other institutes of the National Institutes of Health (multiple grants); the Foundation Fighting Blindness; the American Health Assistance Foundation; PXE International; and Mr. Joseph Cohen. Few things are created without the help of many people. We all stand on the shoulders of our teachers. I particularly acknowledge Drs. Virginia Weimar, Ludwig von Sallmann, and Lorenz E. Zimmerman for the profound influence they have exerted on my career. Producing this book would not have been possible without the full resources of The Jackson Laboratory and the support of its director, Dr. Kenneth Paigen. We are particularly grateful for the efforts of Dr. Barbara Knowles, Beverly Richards-Smith, and Melissa Berry who reviewed all the chapters and provided invaluable information on genetic nomenclature and style. The efforts of the staff of the Joan Staats Library, especially Irene Whitney, are most appreciated; without their patience and perseverance, gathering the hundreds of pieces of reference material would have been most difficult. Adriana Zabeleta and the staff of the Biological Imaging Service are responsible for developing new techniques and for the great care necessary to prepare outstanding histological preparations. We wish to thank Joyce Worcester for graphics assistance and Felicia Farley and Norma Buckley for data entry and editorial assistance. A special debt of gratitude is owed to Drs. John P. Sundberg, Simon W. M. John, and Patsy Nishina for their teaching and constant support and to Linda A. Smith for always being there! Richard S. Smith

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Prefacethe Editors About Richard S. Smith, Allelochemical interactions, M.D., D.during Med. the Sci.,last is three a redecades, search scientist have evolved at The Jackson as an important Laboratory branch in Bar of plant ecology. Harbor, Maine.In Dr. this Smith book, inisgeneral, a 1956 the graduate effects of chemical compounds Wesleyan University with released a B.S. from degree plantswith (includhigh ing microorganisms), honors. He received onhisother M.D. plants from in their Columbia vicinity are considered University Collegeunder of Physicians the term “allelopathy.” and SurgeonsThe in term “allelochemical” 1960, interned at Bellevue is used Hospital, in a wider and returned context in to the Edward field of S. ecology Harkness where EyeitInstitute includes, at but Columbia is not limited for additional to, plant research and microorganism experience andinteractions. a residency Allelochemicals in clinical ophthalmology. released He from received plants a(including Doctor of microorganisms) Medical Science have in Anatomy multifaceted from influences Columbia on ecosystems;in these University 1965. This also was influence followed soilbymicrobial a fellowecology, ship at the soil Armed nutrients, Forces andInstitute physical, of chemical Pathologyand in biological soil ophthalmic pathology factors. and We believe a 2-yearthat appointment it is extraorat dinarily the National difficult Institutes to separate of Health the influence as a research of alleloassochemicals ciate in experimental on each ofocular thesepathology. components Dr. Smith of an ecosystem.a Effects became full-time on any staffonemember of these of compothe nents, due to Department Ophthalmology allelochemicals, at Albany may influence Medical growth,(New Center distribution, York) in and 1968 survival as a of clinical plant species. ophthalmologist Theand aimasofhead this book of theisOphthalmic to provide insight Pathology and recent progress Laboratory. He onwas allelochemical appointed research Professorfrom of this multifaceted Pathology and served standpoint. for 10 Research years as professor articles—reand porting results chairman of theofOphthalmology substantially completed Department. work, He and review came to Thearticles—presenting Jackson Laboratory on novel sabbatical and critical leave appraisals in 1991 andof began specific full-time topics work of there interest, in 1994. are His included. research current Yet it may interests not be include a comprehensive identifyingtreatise mouse on models the subject. of glaucoma, The sequence cataracts,ofand chapters corneal in and the book starts retinal disease, withand ancorrelating overview followed them withbyhuman 34 chapters ocular disease. contributed by scientists around the world, thus presenting a global perspective on allelochemical research. Section I—Methodologies (Chapters 2–8), discusses important aspects of methodology in the study of allelopathy, shortcomings of bioassays for allelopathy, bioassays for different plant groups, extraction of allelochemicals from soil, sampling procedures, and an outline of analytical methods for different classes of allelochemicals. Section II—Interactions Among Plant and Microbial Systems (Chapter 9–15), presents allelochemical research in aquatic and terrestrial ecosystems, and includes other important subjects like pollen allelopathy. Section III—Ecological Aspects (Chapters 16–22), illustrates the significance of ecological studies in allelochemical research, and discusses the important role that the soil environment plays in the functioning of allelochemicals. Section IV—Biochemical, Chemical and Physiological Aspects (Chapters 23–30), discusses biochemical, molecular, and physiological aspects of allelopathy, including information on modes of action of allelochemicals in allelopathy. Allelochemicals have been successfully used in biocontrol of plant pathogens and weeds. This important applied aspect of allelochemistry is discussed under Section V—Biological Control of Plant Disease and Weeds: Applied Aspects (Chapters 31–34). Thus, in totality, the book illustrates the processes, procedures, and applications related to allelochemicals.

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Simon W. M. John, Ph.D., is a staff scientist at The Jackson Laboratory. He received a B.Sc. with joint highest honors in zoology and genetics from University College Cardiff, South Wales in 1985. He completed his Ph.D. in biology at McGill University in 1992 and spent 4 years as a postdoctoral fellow in the laboratory of Dr. Oliver Smithies at the University of North Carolina. He became an associate staff scientist at The Jackson Laboratory in 1995 and an assistant investigator at the Howard Hughes Medical Institute in 1998. He also holds a research assistant professorship in ophthalmology at Tufts University School of Medicine and is a member of the graduate faculty at the University of Maine. His research interests involve the molecular genetics of glaucoma and the control of intraocular pressure. Patsy M. Nishina, Ph.D., is a staff scientist at The Jackson Laboratory. She graduated from the University of Hawaii in 1982 with a B.Ed./B.S. in education and nutrition. She received her master’s and Ph.D. degrees from the University of California, Davis in 1985 and 1988, respectively, and worked as a research and teaching assistant at Davis. She completed a 3-year postdoctoral fellowship at Children’s Hospital in Oakland, California in 1990 and joined the staff of The Jackson Laboratory in 1991. Her primary interest is in mapping genes responsible for retinal degeneration models in mice and in determining their molecular mechanisms. She has also done extensive work on the human genetics of retinal degenerations. In addition, she collaborates with her husband, Dr. Juergen Naggert (a contributing author), on molecular and mapping studies of genes that control obesity and diabetes. John P. Sundberg, D.V.M., Ph.D., is a senior staff scientist at The Jackson Laboratory. Dr. Sundberg graduated from the University of Vermont in 1973 with a B.S. in animal science (summa cum laude) and obtained his D.V.M. degree from Purdue University School of Veterinary Medicine. In 1981, he earned his Ph.D. in comparative pathology and virology from the University of Connecticut. Dr. Sundberg served as assistant professor at the University of Illinois College of Veterinary Medicine from 1981 to 1986, and then joined the staff of The Jackson Laboratory. Dr. Sundberg’s current research interests relate to mouse mutations as models for human and domestic animal dermatological diseases, the comparative pathology and molecular evolution of papillomaviruses, and spontaneous diseases of inbred laboratory mice.

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Contributors Lesley S. Bechtold, M.S. The Jackson Laboratory Bar Harbor, Maine

Gregory Martin, M.S. The Jackson Laboratory Bar Harbor, Maine

Bo Chang, M.D. The Jackson Laboratory Bar Harbor, Maine

James Miller The Jackson Laboratory Bar Harbor, Maine

Norman L. Hawes The Jackson Laboratory Bar Harbor, Maine

Juergen Naggert, Ph.D. The Jackson Laboratory Bar Harbor, Maine

John R. Heckenlively, M.D. Jules Stein Eye Institute University of California Los Angeles, California

Patsy M. Nishina, Ph.D. The Jackson Laboratory Bar Harbor, Maine

Sakae Ikeda, Ph.D. The Jackson Laboratory Bar Harbor, Maine Simon W. M. John, Ph.D. The Jackson Laboratory Bar Harbor, Maine Winston W.-Y. Kao, Ph.D. Department of Ophthalmology University of Cincinnati Cincinnati, Ohio Carol C. Linder, Ph.D. The Jackson Laboratory Bar Harbor, Maine Chia-Yang Liu, Ph.D. Department of Ophthalmology University of Cincinnati Cincinnati, Ohio

Steven Nusinowitz, Ph.D. Department of Ophthalmology Jules Stein Eye Institute University of California Los Angeles, California Timothy O’Brien, Ph.D. The Jackson Laboratory Bar Harbor, Maine Melissa J. Relyea, B.S. The Jackson Laboratory Bar Harbor, Maine William H. Ridder III, O.D., Ph.D. Southern California College of Optometry Fullerton, California Olga V. Savinova, M.S. The Jackson Laboratory Bar Harbor, Maine

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Richard S. Smith, M.D., D. Med. Sci. The Jackson Laboratory Bar Harbor, Maine John P. Sundberg, D.V.M., Ph.D. The Jackson Laboratory Bar Harbor, Maine

Adriana Zabaleta, M.S. The Jackson Laboratory Bar Harbor, Maine

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Table of Contents SECTION I

Regional Anatomy and Development

Chapter 1 The Anterior Segment and Ocular Adnexae ......................................................................................3 Richard S. Smith, John P. Sundberg, and Simon W. M. John Chapter 2 The Posterior Segment and Orbit ....................................................................................................25 Richard S. Smith, Simon W. M. John, and Patsy M. Nishina Chapter 3 Ocular Development........................................................................................................................45 Richard S. Smith, Winston W.-Y. Kao, and Simon W. M. John

SECTION II

Issues in Mutant Mouse Evaluation

Chapter 4 Strain Background Disease Characteristics.....................................................................................67 Richard S. Smith and John P. Sundberg Chapter 5 Selection of Controls .......................................................................................................................77 John P. Sundberg, Richard S. Smith, and Simon W. M. John Chapter 6 Genetic Approaches to Identifying QTL and Modifiers of Hereditary Ocular Phenotypes.......................................................................................................................................81 Juergen Naggert and Patsy M. Nishina Chapter 7 Mutagenesis and Genetic Screens in the Mouse .............................................................................93 Timothy O’Brien

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SECTION III Regional Ocular Pathology Chapter 8 The Anterior Segment....................................................................................................................111 Richard S. Smith, John P. Sundberg, and Simon W. M. John Chapter 9 Choroid, Lens, and Vitreous ..........................................................................................................161 Richard S. Smith Chapter 10 Retina.............................................................................................................................................195 Richard S. Smith, Norman L. Hawes, Bo Chang, and Patsy M. Nishina Chapter 11 Optic Nerve and Orbit ...................................................................................................................227 Richard S. Smith, Simon W. M. John, and John P. Sundberg

SECTION IV Biomethods Chapter 12 Photography and Necropsy............................................................................................................251 Richard S. Smith, Norman L. Hawes, James Miller, John P. Sundberg, and Simon W. M. John Chapter 13 General and Special Histopathology .............................................................................................265 Richard S. Smith, Adriana Zabaleta, Simon W. M. John, Lesley S. Bechtold, Sakae Ikeda, Melissa J. Relyea, John P. Sundberg, Chia-Yang Liu, and Winston W.-Y. Kao A. Light Microscopy.....................................................................................................................266 Richard S. Smith, Adriana Zabaleta, and Simon W. M. John B. Electron Microscopy................................................................................................................272 Lesley S. Bechtold and Richard S. Smith C. Immunohistochemistry ............................................................................................................277 Sakae Ikeda, Melissa J. Relyea, and John P. Sundberg D. Northern and in Situ Hybridization..........................................................................................285 Sakae Ikeda, Chia-Yang Liu, and Winston W.-Y. Kao

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Chapter 14 Other Techniques ...........................................................................................................................299 Gregory Martin, Richard S. Smith, Simon W. M. John, Olga V. Savinova, Steven Nusinowitz, William H. Ridder III, and John R. Heckenlively A. Cell Kinetics, Morphometrics, and Confocal Microscopy ......................................................300 Gregory Martin and Richard S. Smith B. Intraocular Pressure Measurement in Mice: Technical Aspects ..............................................313 Simon W. M. John and Olga V. Savinova C. Electrophysiological Testing of the Mouse Visual System......................................................320 Steven Nusinowitz, William H. Ridder III, and John R. Heckenlively Chapter 15 Resources.......................................................................................................................................345 Carol C. Linder Index .............................................................................................................................................353

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Section I Regional Anatomy and Development

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The Anterior Segment and Ocular Adnexae Richard S. Smith, John P. Sundberg, and Simon W. M. John

CONTENTS Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .3 Lids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .6 Conjunctiva . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .9 Lacrimal System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .9 Cornea and Sclera . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .9 Anterior Uvea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .13 Iris . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .13 Iris Stroma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .13 Iris Pigment Epithelium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .13 Iris–Ciliary Body Junction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .15 Ciliary Body . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .15 Aqueous Drainage System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .17 Lens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .19 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .21

INTRODUCTION Appropriate evaluation of any organ system requires thorough knowledge of gross, microscopic, and ultrastructural features. The first section of this book focuses on demonstrating and discussing this critical information. An understanding of ocular development can enhance comprehension of the relationships of ocular structures as well as the mechanisms that underlie congenital abnormalities. This latter topic is discussed in Chapter 3. Although the eye is often considered a distinct structure or organ, it is surrounded and maintained by adjacent soft and bony tissues, all of which are discussed in this and the following chapter. The anterior segment of the eye, the subject of this chapter, includes the conjunctiva, cornea, anterior chamber, aqueous drainage structures, iris, ciliary body, and lens. The ocular adnexae include the various components of the lids and the lacrimal glands.

LIDS The palpebrae (eyelids) are complex structures, consisting of skin, mucous membrane, smooth and striated muscle, the tarsal plate, and glandular structures. A thin layer of epidermis that consists of 7 to 12 layers of keratinocytes (epithelial cells) covers the haired skin surface, thickens at the mucocutaneous junction at the lid margin, then abruptly changes into the conjunctiva. The sharp transition is best demonstrated by immunohistochemistry. The cutaneous suprabasal epidermis expresses keratins 1 and 10, whereas the conjunctiva expresses keratin 6. Both epidermis and conjunctiva (Figure 1.1) express keratin 14. The basal keratinocytes located at the dermal/epidermal junction have a low cuboidal to columnar shape. The keratinocytes become flattened at the surface of the epidermis and then become cornified. The cutaneous surface of the lids is covered by hair similar to truncal pelage 0-8493-0864-X/02/$0.00+$1.50 © 2002 by CRC Press LLC

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FIGURE 1.1 Lids-scanning electron microscopy. A. Adult mouse eyelid. The boxed area is enlarged in B. B. Mucocutaneous junction of the eyelid. Numerous cilia (arrows) arise from the skin of the lid. Keratin flakes are visible on the epidermal surface between the cilia. Posterior to the mucocutaneous junction, the conjunctiva (C) has a smooth surface. C to F. Cytokeratin staining of the eyelid of a normal C57BL/6J mouse. C. Keratin 1 stains the suprabasal epidermis (E), but not the conjunctiva (C). Positive staining ceases at the mucocutaneous junction (arrow). D. The staining pattern is reversed with keratin 6, with staining of the conjunctiva, but not the epidermis. The staining pattern also demarcates the mucocutaneous junction. E. Keratin 10 staining demonstrates a pattern similar to keratin 1. F. Keratin 14 stains both epidermis and conjunctiva, as well as the epithelial cells of the Meibomian gland. C to F original magnification × 100.

hairs. Cornification disappears at the lid margin. The cilia (eyelashes) are located just anterior to the mucocutaneous junction (Figures 1.1 and 1.2). Posterior to the mucocutaneous junction the epidermis is much thinner, lacks cornification, and becomes the tarsal conjunctiva (Figure 1.2).1,2 The lacrimal punctae are located at the medial aspect of the dorsal and ventral palpebrae and mark the opening of the lacrimal drainage system that ultimately terminates in the nasal cavity. The orifices of the Meibomian glands are located just behind the mucocutaneous junction along the lid margin. Their secretions lubricate the lids and contribute to the tear film.

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FIGURE 1.2 Lids. A. Coronal section through eyelid. The surface epidermis (arrow) has fine folds, whereas the conjunctival surface (arrowhead) is smooth and contains goblet cells. Meibomian glands (M) are located just beneath the conjunctiva. A hair follicle (*) is the source of the cilia that are located at the mucocutaneous junction. Original magnification × 20. B. Secretions from the Meibomian glands (M) enter a duct (*) that opens on the lid surface at the mucocutaneous junction. Original magnification × 50. C. The surface epidermis (arrow) of much of the lid is thin and lightly cornified. The striated muscle of the levator palpebrae (L) is located just beneath the dermis. Original magnification × 100. D. A higher magnification of A demonstrates the presence of goblet cells (arrowhead) some distance from the lid margin. Between this point and the lid margin, the conjunctiva consists of smooth, non-cornified epidermis (*). The bulb of a ciliary hair follicle is indicated by an arrow. Original magnification × 200. E. A few lymphocytes are often seen beneath the conjunctival epithelium (*), but this does not indicate infection. Original magnification × 400. F. Close to the lid margin, where exposure is greatest, the cornified layer of the epidermis (arrowhead) is thickened and the epidermis has prominent rete ridges (arrow). Original magnification × 100.

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The palpebrae are supported by the tarsal plate that is composed of dense collagen and by the striated orbicularis oculi muscle (supplied by cranial nerve VII) that functions to close the eye. The dorsal lid is raised by the striated levator muscle (Figure 1.3), which is closely associated with the superior rectus muscle (see below).3 The orbital contents are surrounded by a membranous smooth muscle known as the orbital muscle. The orbital muscle attaches to the levator muscle as well as to the bony wall of the orbit and may correspond in part to primate periorbita, the ligaments of Lockwood and Whitnall, and the tarsal (Muller’s) muscle.4-6 The cilia are closely associated with large modified sebaceous glands, the Meibomian glands, that open just posterior to the cilia along the mucocutaneous junction. Small sebaceous glands (glands of Zeis) are attached to the cilial follicles. Information concerning the nerves and vascular supply of lid and orbital structures of mice is limited. It is likely that the trigeminal (V) nerve provides the sensory supply to the eye and orbit, that the facial nerve (VII) innervates the orbicularis muscle, and that the oculomotor nerve (III) supplies the levator muscle.7,8 Adrenergic fibers from the superior cervical ganglion and cholinergic fibers from the pterygopalatine ganglion have been traced to the tarsal smooth muscle in mice.6 Their exact anatomic relationships and variations remain unknown. The same limitations apply to detailed knowledge of the vascular supply of the lid that is supplied by the external carotid and drained via the maxillary vein to the external jugular vein. Superior and inferior orbital arterial and venous branches have been briefly described.7,8 Aging changes. Ulcerative blepharitis (Figure 1.3) is a lesion often identified in susceptible strains, including 129/J, BALB/cJ, BALB/cByJ, CBA/J, and A/HeJ.9,10 Both the epidermis and accessory glands of the lids may be involved in the process and periorbital abscess may be an accompanying lesion. Ulcerative blepharitis typically becomes more severe with age and likely has a genetic basis since it occurs at a higher rate in specific inbred strains (see also Chapters 4 and 8).9,10

CONJUNCTIVA The conjunctiva begins at the mucocutaneous junction and continues over the inner surface of the palpebrae to the dorsal and ventral fornix (tarsal conjunctiva), where it is reflected back (fornix) to cover the sclera, extending to the corneoscleral junction (bulbar conjunctiva). Rostrally, there are dorsal and ventral folds of conjunctiva. The dorsal fold of conjunctiva forms the nictitating membrane (Figure 1.4), supported by a small plate of cartilage. The ventral fold contains only loose connective tissue.11 The bulbar conjunctival surface is composed of two to four layers of keratinocytes that originate from a cuboidal basal layer. The tarsal conjunctiva consists of four to seven cell layers. The conjunctival keratinocytes are flattened toward the free surface, but do not normally become cornified. In addition to the keratinocytes, numerous mucin-secreting goblet cells are spread over the tarsal and bulbar conjunctival surfaces (see Figure 1.3). The goblet cells are easily visualized with alcian blue, mucicarmine, or periodic acid–Schiff (PAS) stains. The fornical epithelium consists of only one to two cell layers and is the primary source of stem cells for both keratinocytes and goblet cells, although a few stem cells are scattered throughout the conjunctiva.12 Although yet to be demonstrated in the mouse, the primary source for replacement of conjunctival epithelial cells in rabbits is the mucocutaneous junction.13 Beneath the keratinocytes of the conjunctiva, there is a thin layer of loose, heavily vascularized, collagenous connective tissue, lightly attached to the underlying sclera. Lymphocytes, eosinophils, and mast cells are commonly found in this location.14,15 In pigmented strains, there may be foci of pigment-filled macrophages beneath the conjunctiva (pigmentary incontinence). This is almost a constant finding in the conjunctiva that covers the nictitating membrane.

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FIGURE 1.3 Lids. A. The conjunctiva is very thin (*) in the conjunctival fornix, where the conjunctiva covering the lid is reflected onto the sclera (S). A thin layer of connective tissue (arrow) separates the conjunctiva from the sclera and from the levator (L) muscle. Original magnification × 200. B. Goblet cells are particularly abundant (arrows) in the depths of the fornix. Original magnification × 400. C. The conjunctival epithelium (E) consists of four to five layers of cells. Dermal collagen separates the epithelium from a fibroblast (F). A goblet cell (*) has discharged its contents into the tear film. Original magnification × 15,000. D. In many places the individual cells of the conjunctival epithelium are extremely thin and contain electron-dense opacities (arrowheads) of unknown composition. The most superficial epithelial cells have many microvilli (arrows) on the surface that are said to help maintain the integrity of the tear film. Original magnification × 20,100. E. In a CBA/J mouse, there is a large focus of ulcerative blepharitis (*). Original magnification × 100. F. At higher magnification, the epidermis is absent between the arrows. There is a large focus of coagulative necrosis and chronic inflammation (*). Original magnification × 200.

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FIGURE 1.4 Lacrimal and cornea. A. The lacrimal gland (L) is small and located in the anterior orbit. Its posterior aspect is indented by the Harderian (H) gland. Original magnification × 50. B. The lacrimal gland secretory cells are arranged in acini (arrow). Tears are carried to the ocular surface by small ducts lined by low cuboidal epithelium arrowheads. Original magnification × 630. C. The corneal epithelium (E) is normally five to six layers thick. Mitosis is frequently observed (arrowhead) and generally limited to the basal layer. Bowman’s layer (arrow) is acellular. The collagen of the corneal stroma (S) is arranged in orderly layers parallel to the surface and the keratocytes are flattened in a similar orientation. Descemet’s membrane (*) separates the stroma from the corneal endothelium (open arrow). Original magnification × 630. D. At the limbus, the cornea undergoes a rapid transition to the sclera. The corneal epithelium becomes the much thinner conjunctival epithelium (E). A thin layer of loose connective tissue (*) lies between epithelium and sclera. Note the irregular arrangement of scleral collagen compared with the corneal stroma in C. Vascular channels (arrow) are absent from the cornea, but abundant in the sclera at the limbus. Original magnification × 400. E. The nictitating membrane (arrow) is located by the rostral end of the lid fissure. Original magnification × 40. F. The shape of the long nictitating membrane is maintained by a thin band of cartilage (arrow) that is covered by loose connective tissue and by conjunctival epithelium. Original magnification × 100.

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LACRIMAL SYSTEM Mice have two pairs of lacrimal glands that empty into the conjunctival sac by a common duct (see Figure 1.4). The orbital gland is small and located laterally beneath the upper lid. The extraorbital lacrimal gland is located anterior and ventral to the ear, adjacent to the parotid gland. Both lacrimal glands are surrounded by a capsule and divided by connective tissue into lobules. The individual alveoli are larger and less densely packed than those of the parotid gland. The cuboidal or low columnar cells are characterized by granular basophilic cytoplasm. The nuclei have a basal location in the glandular epithelium. Specialized myoepithelial cells surround both acini and excretory ducts.16 A single layer of cuboidal epithelium lines the intralobular and excretory ducts (Figure 1.4).8,17 The combination of aqueous tears from the lacrimal glands, mucin from the goblet cells of the conjunctiva, and the oily Meibomian secretion make up the complex tear film that protects the corneal and conjunctival surfaces from drying, from microtrauma by foreign bodies, and from bacterial invasion.18,19 In primates, the tear fluid is drained from the eye through the nasal dorsal and ventral lacrimal punctae into the lacrimal sac via the lacrimal canaliculi and from the lacrimal sac into the nose through the nasolacrimal duct. A similar pathway is followed in mice.20

CORNEA AND SCLERA The cornea is composed of five layers: epithelium, Bowman’s layer, stroma, Descemet’s membrane, and endothelium (Figures 1.4 and 1.5).21 The keratinocytes of the corneal epithelium are similar in appearance to those of the conjunctiva, except goblet cells are absent. Under normal circumstances, the corneal keratinocytes are not cornified, although they do become flattened as they approach the surface. Mitosis is limited to the low columnar basal cells that are attached to a thin basal lamina by hemidesmosomes. Desmosomal attachments between adjacent keratinocytes are prominent in all but the most superficial cells. Electron microscopy demonstrates complex interdigitations between adjacent keratinocytes.22 In primates, Bowman’s layer is easily visualized and consists of a random fibrillar network of type I collagen fibrils23 and may also contain type XII collagen during development.24 The subepithelial collagen fibers in mice are also arranged in random fashion, corresponding to a thin Bowman’s layer, although some believe mice do not have a true Bowman’s layer.25,26 It may be difficult to visualize Bowman’s layer in light microscopic preparations, but it is clearly demonstrable by electron microscopy.27 Clear vision in all vertebrates is dependent on transparency of the cornea, particularly the corneal stroma. The cornea is transparent for three reasons: (1) all corneal collagen fibers have a constant uniform diameter; (2) they are regularly spaced with regard to each other and bundles of fibers are also regularly arranged, parallel to the corneal surface; and (3) the cornea is relatively dehydrated compared with other connective tissue such as the sclera. Mouse stromal collagen fibers have a diameter of 29 ± 4 nm and maintain a center-to-center distance of 55 ± 100 nm.27 The corneal stroma accounts for two thirds of the total corneal thickness. The keratocytes (a term not to be confused with keratinocytes, the cells of the epidermis) are modified fibroblasts with elongated oval nuclei and are arranged parallel to the collagen bundles. Keratocyte cytoplasm is scanty and contains abundant endoplasmic reticulum. The normal mouse cornea is completely avascular, another feature important for corneal transparency. Sensory innervation of the cornea is supplied by axons from the trigeminal nerve.28 The posterior termination of the corneal stroma is bordered by Descemet’s membrane, a highly specialized basal lamina (see Figure 1.5) that is produced by the corneal endothelium. Descemet’s membrane on the stromal side has a highly ordered hexagonal array of very fine fibers composed of laminin, fibronectin, and type IV collagen and often has a banded appearance.23,26 The anterior portion

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FIGURE 1.5 Cornea. A. The cuboidal basal cells (B) of the cornea are the source of the more superficial cells. The corneal epithelial cells are held together by desmosomes and adherent junctions (arrowheads). As the cells approach the surface they become flattened. The most superficial cells have numerous microvilli (arrow). Asterisk indicates Bowman’s membrane. Original magnification × 20,100. B. The collagen of the corneal stroma is arranged in parallel bundles, oriented with their long axis to the surface of the eye. The mild distortion of the collagenous lamellae in this image is due to a slightly tangential plane of section. The keratocytes (arrow) are flattened and have little cytoplasm. Original magnification × 21,100. C. The corneal stroma (S) is bordered internally by Descemet’s membrane (D). Descemet’s membrane is dense externally. Internally, long-spacing collagen (arrowhead) is abundant. The corneal endothelium (arrow) faces the anterior chamber. Original magnification × 15,000. D. The vigorous metabolic activity of the corneal endothelium is indicated by the abundant mitochondria (arrowhead) and endoplasmic reticulum (arrow). The boxed area is enlarged in the inset to show the location of the tight and gap junctions that seal the extracellular space between adjacent cells. Original magnification × 12,000; inset × 30,000.

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of Descemet’s membrane is produced before birth. On the anterior chamber side, Descemet’s membrane is more granular and homogeneous, resembling typical basal lamina; it becomes thicker with age. Peripherally, especially in mice over two months of age, short segments of “long-spacing” 100nm collagen fibrils are often present (see Figure 1.5). Descemet’s membrane terminates at the corneoscleral junction, just anterior to the trabecular meshwork. In most instances this termination is gradual in mice, but a focal thickening may be observed in some strains that corresponds to Schwalbe’s line in primate eyes.29 The posterior aspect of Descemet’s membrane is covered by a layer of cells commonly called endothelial cells, although they most closely resemble mesothelium and are of nonvascular origin. This layer of endothelial cells is critical for corneal transparency since the cells transport fluid out of the stroma, maintaining corneal dehydration. The role of the endothelium in active electrolyte transport is suggested by the abundance of mitochondria and pinocytotic vesicles. Adjacent endothelial cells are attached by both loose and gap junctions.23,26 Primate endothelial cells do not undergo significant mitosis after birth.30 The status of mouse endothelial cells in this regard is unknown, although bromodeoxyuridine (BrdU) studies conducted over a 3-month period failed to show any BrdU-staining endothelial cells after birth (Smith and Nishina, unpublished observations). The cornea blends into the sclera at the corneoscleral junction (limbus) (see Figures 1.4 and 1.5). It is generally accepted that the stem cells that renew the corneal epithelium after injury are located at the limbus.31,32 Langerhans cells are present at the limbus and beneath the epithelium in the peripheral cornea. These cells are similar to those found in the epidermis and they express HLA-DR (Ia) antigens in human eyes (H2 antigens for mice).33 Ocular Langerhans cells are not evident with routine histological stains, but stain positively with ATPase as well as with specific antibodies to HLA-DR (human) or H2 (mouse) antigen.34 These cells are present in mice and other mammals,35,36 and play an important role in graft rejection and in inflammatory reactions to exogenous antigens.37,38 Posterior to the limbus, the sclera is covered by bulbar conjunctiva until the fornix is reached. Posterior to that point, the sclera is covered by loose connective tissue and other orbital structures (see Chapter 2). At the limbus, the highly ordered bundles of corneal collagen give way to a random arrangement of collagen that results in a layer slightly less thick than the cornea. The sclera is thickest posteriorly and in the mouse is so thin anteriorly that the underlying uvea is easily visualized in pigmented mice. The uniform diameter of corneal collagen fibers disappears, and scleral collagen has a broad range of fibril diameters. In addition, blood vessels are present posterior to the limbus and form a limbal arcade encircling the cornea, as occurs in other vertebrates. Although studies have not been done in mice, the rat anterior segment has a circumferential vascular ring and venous plexus that is supplied by the anterior and long posterior ciliary artery system.39 Rarely, episcleral nerve loops from the long posterior ciliary nerves penetrate the sclera just posterior to the limbus. In a pigmented mouse, this often results in a pigmented subconjunctival spot (Figure1.6A). Aging Changes—Cornea. In some strains, corneal subepithelial and anterior stromal mineralization are common. Although not limited to these strains, it is particularly frequent in DBA/2J, as well as some C3H and BALB/c substrains, and in both AKXD and BXD recombinant inbred strains. Most of these same strains are prone to cardiac calcinosis, presumably secondary to metastatic mineralization.40 The corneal deposits are extracellular, basophilic, often have a globular appearance, and may be associated with thinning or ulceration of the overlying corneal epithelium (Figure 1.6B). The deposits often thicken and increase in diameter with age. The mineral nature of the deposits can be confirmed with von Kossa stain. Corneal neovascularization and keratitis may occur in older mice. The occasional observation of foreign material suggests that some of this may be related to trauma from bedding or from other mice in the cage. Strains prone to blepharoconjunctivitis are also at increased risk for keratitis and its sequelae.

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FIGURE 1.6 Cornea and anterior chamber. A. An episcleral nerve loop (arrow) is formed when a long posterior ciliary nerve extends into the limbus. These often carry pigment with them. Original magnification × 100. B. Platelike deposits of calcium and other minerals (arrows) are seen in older mice of many strains. Original magnification × 200. C. The anterior chamber angle (*) is located at the junction between the cornea (C) and iris (arrow). Ciliary processes (CB) are located on the peripheral posterior aspect of the iris and over the ciliary muscle. The aqueous drainage structures include the trabecular meshwork (arrowhead) and Schlemm’s canal (open arrow). Original magnification × 100. D. The entrance into the trabecular meshwork (arrow) is defined by the deep angle recess (*). Schlemm’s canal (SC) extends from a point by the end of Descemet’s membrane to a position external to the posterior aspect of the ciliary body (CB). Original magnification × 630. E. An iris process (arrow) extending from the iris surface to the peripheral cornea is a common, normal finding. Original magnification × 200. F. Schlemm’s canal (*) connects directly to the aqueous veins (arrows) that in turn connect to the systemic circulation. Methylene blue. Original magnification × 630.

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ANTERIOR UVEA IRIS The iris separates the anterior and posterior chambers and is the most anterior part of the uvea. The iris sphincter and dilator muscles work together to control pupillary diameter, providing a variable diaphragm for the eye’s optical system and controlling the amount of light that reaches the retina. The iris and ciliary body are attached to the anterior sclera at the iris root, and the anterior chamber angle is formed at the point where the iris and cornea meet. The aqueous outflow pathways (see below) also lie adjacent to the iris root. In the mouse eye, the pupillary border of the iris is usually in contact with the anterior lens capsule. Space exists between these two structures, because the aqueous humor, formed by the ciliary body, flows freely into the anterior chamber through the pupil.

IRIS STROMA The iris consists of the stroma and two-layered pigment epithelium. The stroma is a derivative of cranial paraxial mesoderm and neural crest. With light microscopy, much of the thin iris stroma appears empty but such spaces actually contain dispersed collagen and components of the extracellular matrix (Figures 1.6 and 1.7). The anterior surface of the iris is often referred to as the anterior border layer.41 However, this term is a misnomer, since there is no delimiting membrane or series of junctional complexes that separate the iris stroma from the anterior chamber.42 Small blood vessels found in the stroma arise from the anterior branches of the ciliary arteries and are surrounded by a discontinuous pericyte layer. These vessels are thick walled, presumably to prevent injury from the motions of the iris as it dilates and constricts. Tight junctional complexes seal the vascular endothelium and make them impermeable to small tracers such as horseradish peroxidase. 42,43 Two types of pigmented cells are found in the stroma: dendritic melanocytes and iris clump cells. The dendritic melanocytes contain numerous melanosomes. The content of melanosomes and the extent of their pigmentation determine iris color. In albino mice, the melanosomes are still present, but lack melanin. The iris clump cells are presumably macrophages that have ingested free melanosomes. They are rounded cells most frequently found in the iris root and near the pupil. The number found varies from strain to strain. In some strains of mice (see Section III), extensive accumulations of these pigmented macrophages are observed. Fibroblasts have been reported in human iris,41 but they are not abundant. Fibroblasts have not been demonstrated in mouse iris. Typical myelinated nerves are easily identified in the iris stroma and unmyelinated nerves are also present. Although not apparent by routine light microscopy, near the anterior iris surface, there are abundant dendritiform macrophages and MHC class II+ dendritic cells that are antigen-presenting cells. These cells can only be identified by appropriate immunohistochemical techniques.44

IRIS PIGMENT EPITHELIUM The iris pigment epithelium and the sphincter and dilator muscle arise from the anterior portion of the optic cup (neuroepithelium). For developmental reasons (see Chapter 3) the apices of the epithelial cells are apposed. As a result, the basal lamina of the posterior iris pigment epithelium faces the vitreous while the anterior epithelial basal lamina faces the stroma. The lateral and apical faces of the cells demonstrate extensive interdigitations maintained by desmosomal and other loose junctional complexes. The cytoplasm of the pigment epithelium contains abundant melanosomes that are more rounded than those found in the anterior stroma. The anterior layer of iris pigment epithelium is the source of both the iris sphincter and dilator smooth muscles. They represent one of the few situations in which muscle tissue is derived from neuroepithelial rather than mesenchymal sources. Although the smooth muscle arises directly from the epithelial cells, it is ultrastructurally identical to smooth

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FIGURE 1.7 Iris and pars plana. A. The pars plana (arrows) lies between the ciliary body (CB) and retina (R). Original magnification × 100. B. The pars plana (arrows) consists of a single layer of pigmented epithelium that is continuous with the retinal pigment epithelium and a single layer of nonpigmented epithelium that becomes the retina. Original magnification × 400. C. The anterior iris stroma (S) contains blood vessels and many dendritic melanocytes. The bilayered posterior iris pigment epithelium (arrow) is separated from the stroma by the thin dilator muscle (arrowhead). Original magnification × 630. D. The iris sphincter muscle (*) is located adjacent to the pupillary border. Original magnification × 630. E. The iris stroma contains many dendritic melanocytes (arrow), blood vessels (V), and nerves (N). Although the anterior border of the iris appears continuous at some levels of section, there are points (arrow, inset) where the iris stroma is open to the anterior chamber. Original magnification × 12,000. F. The posterior iris pigment epithelium (arrow) is filled with melanosomes. The anterior iris pigment epithelium (open arrow) is thinner and is the source of the delicate smooth muscle of the iris dilator (*). Original magnification × 8100. G. Near the pupillary border, the posterior pigment epithelium (arrow) becomes attenuated and extends no farther than the pupillary border. The most prominent structure in this location is the smooth muscle of the iris sphincter (M). Original magnification × 8100.

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muscle from other sources (see Figure 1.7), with basal lamina, pinocytotic vesicles, focal plasma membrane densities, and cytoplasmic actin and myosin. The two layers of iris pigment epithelium are continuous with the ciliary body epithelium.

IRIS–CILIARY BODY JUNCTION The ciliary body is directly attached to the overlying sclera in human eyes.45 The arrangement in mice is different in that the stroma of the iris merges without obvious boundary into the stroma of the ciliary body. In turn, the most anterior ciliary processes arise from the posterior aspect of the peripheral iris and, in mice, only the posterior ciliary processes attach directly to the sclera. Thus, some of the external face of the ciliary body is almost in contact with the posterior trabecular meshwork. Over the posterior aspect of the peripheral iris and extending back over the adjacent sclera, the ciliary epithelium with its vascular supply are prominently folded to form the ciliary processes.

CILIARY BODY Functionally, the ciliary body produces both the aqueous humor and some components of the vitreous and is the source of the zonules that support the lens. The ciliary body (Figure 1.8) is composed of three layers: epithelium, vascular stroma, and muscle. The external layer of ciliary epithelium is pigmented and continuous with the retinal pigment epithelium and the anterior iris pigment epithelium. The internal (facing the vitreous) layer of ciliary epithelium is nonpigmented and continuous with the retina. Each epithelial layer has a different function. The apices of the pigmented (PE) and nonpigmented ciliary epithelium (NPE) are apposed, as they are in the iris pigment epithelium. The base of the NPE faces the vitreous, whereas the base of the PE faces the vascular stroma. The zonules, which are also attached to the lens capsule (see below), are embedded in the basal lamina and plasma membrane of the NPE of the ciliary processes. Special stains for glycosaminoglycans such as colloidal iron or alcian blue are strongly positive in the NPE, an indication of the production of adult vitreous by the NPE.46-48 The lateral cell membranes of both layers are held together primarily by open junctions (zonulae adhaerentes), although short tight junctions are often found between adjacent cells near the cell apex. The apical extracellular space between PE and NPE layers is much more complex, reflecting the functions of the ciliary epithelium in production of the aqueous humor. For the most part, the space is closed by zonulae adhaerentes or by typical gap junctions. However, the extracellular space has periodic open regions partly filled with villous cytoplasmic processes, known as ciliary channels.42,43 In the cytoplasm of NPE cells, there are abundant mitochondria, endoplasmic reticulum, and a prominent Golgi apparatus that reflect a high level of cell activity. Pinocytotic vesicles are also prominent and transport low-molecular-weight tracer substances, such as horseradish peroxidase.42,43 This finding is consistent with the selective barrier function of the ciliary epithelium in aqueous humor formation and the presence of small amounts of protein in the aqueous. Although not apparent without appropriate immunohistochemistry, there are MHC class II+ dendritic cells within the NPE. 44 The ciliary body PE has a basal lamina that merges with the basal lamina of the immediately adjacent ciliary capillaries. The base of the PE cell is characterized by complex infoldings of the plasma membrane. Laterally, near the apex of the PE cell, adherent junctions join adjacent cells, but elsewhere the extracellular space that surrounds the PE cell is open between adjacent cells. In the cytoplasm, there are numerous large mitochondria and, in pigmented mice, many round melanosomes. The plasma membranes of the cell apex contribute to the complex structure of the apical extracellular space along with the NPE cells.42,43 The vascular supply of the ciliary body is derived from branches of the long posterior ciliary arteries. The small capillary channels that lie in contact with the PE are fenestrated capillaries that allow rapid transport of materials in and out of the adjacent PE cells as the aqueous humor (a selective

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FIGURE 1.8 Ciliary body. A. The ciliary body includes the vascular stroma and ciliary muscle (arrow) and the ciliary processes (CP). Original magnification × 630. B. The vitreal surface of the ciliary processes is covered by nonpigmented ciliary epithelium (NPE, arrowhead), beneath which is the pigmented ciliary epithelium (arrow) and vascular channels. Original magnification × 630. C. The basal lamina (arrowhead) of the nonpigmented ciliary epithelium (N) faces the posterior chamber and vitreous (V). The apical extracellular space of the NPE is sealed by short tight junctions (arrow). In places, the space between NPE and pigmented ciliary epithelium (PE) the extracellular space is widened and filled with membranous processes (open arrow), the ciliary channels. The ciliary capillaries (*) are fenestrated and abut directly on the base of the PE cells and their elaborately infolded basal cell membranes. Original magnification × 12,000.

ultrafiltrate) is formed. Small amounts of connective tissue separate the capillary bed from the ciliary muscle. In primates, the ciliary muscle is substantial, with a triangular shape.45 In mice, the ciliary muscle is small and has a cylindrical shape. The smaller muscle is consistent with the lack of accommodation (focusing) in mice. The first smooth muscle fibers can be identified just anterior to the

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termination of the retina and normally extend forward to a point just anterior to the posterior termination of Schlemm’s canal. The muscle is demarcated internally by the capillary bed and externally by the sclera. Muscle fibers may surround the posterior end of Schlemm’s canal, although there is considerable variation in different inbred strains. Typical dendritic melanocytes are evident throughout the stroma of the ciliary body in pigmented mice.

AQUEOUS DRAINAGE SYSTEM In all vertebrates, the clear aqueous humor is formed by the ciliary body, passes through the pupil into the anterior chamber (the area bounded by iris, pupil, and posterior corneal surface), and then leaves the eye through the aqueous drainage system. The balance between secretion and drainage determines constant intraocular pressure. The basic elements of this system are the trabecular meshwork, Schlemm’s canal, the uveoscleral meshwork, and the aqueous veins (Figures 1.6, 1.7, and 1.9). Since corneal stroma, Descemet’s membrane, corneal endothelium, and the trabecular meshwork are all derivatives of a mixture of neural crest49 and cranial paraxial mesoderm50 that begin their formation early in development of the eye (see Chapter 3), it is not surprising that their structures are essentially continuous. The trabecular meshwork consists of a series of trabecular beams that begins just posterior to the termination of Descemet’s membrane and continues posteriorly to a point midway over the ciliary body. There are three to four anterior and seven to ten posterior trabecular beams in mice. This represents about one third to one half the number in the human eye (see Figure 1.9). At some levels of section, the trabecular meshwork appears cut off from the anterior chamber by thick iris processes (see Figure 1.6) that are attached to the slightly enlarged termination of Descemet’s membrane known as Schwalbe’s line. Such iris processes are frequently identified in mice. The “cutoff” area has been referred to as the “spaces of Fontana.” However, serial sectioning in mice demonstrates that the iris processes are not continuous. If they were, then aqueous circulation would be impossible.51 In primates and mice, trabecular beams exhibit a similar architecture. There is a central core of collagen oriented parallel with the long axis of the beam. This is intermixed with scattered small deposits of elastic tissue best seen by electron microscopy. The core is surrounded on all sides by basal lamina material resembling the inner aspect of Descemet’s membrane. Elongated, attenuated endothelial cells cover the outside of each beam. The thickness of trabecular beams is slightly greater in the posterior meshwork. Adjacent beams are usually separated by a space about the same width as the beams, except in the area directly adjacent to Schlemm’s canal (see Figure 1.9).51 The intertrabecular spaces are quite narrow and the extracellular matrix dense, a feature that corresponds to what is termed the juxtacanalicular connective tissue in primates.29,52,53 In primate eyes the trabecular meshwork is divided into corneoscleral (anterior) and uveoscleral (posterior) meshwork. The mouse eye has similar architectural features with a greater number of tightly packed beams in the posterior meshwork. Some feel that there is significant flow of aqueous humor through the uveoscleral route (in animals other than mice), whereas others do not.52-55 The relative contribution of drainage through the uveoscleral route vs. outflow through Schlemm’s canal has not been investigated in mice. Schlemm’s canal is a circumferential, endothelial-lined channel that lies just exterior to the trabecular meshwork. It is continuous around the limbus, but is often briefly and locally divided by collagenous bridges that are covered by endothelial cells. Schlemm’s canal connects directly to the aqueous veins that continue to the scleral surface, where they drain through the venous drainage pathways of the conjunctiva and orbit (see Figures 1.6 and 1.9). Careful examination of the inner wall of Schlemm’s canal reveals giant vacuoles within endothelial cells. These are best seen by electron microscopy where serial sections can demonstrate a passage from one side of the cell to the other.56 In normal mice, several layers of smooth muscle that arise from the ciliary body mark the posterior termination of Schlemm’s canal. Unlike the primate eye, there is no scleral spur in this region.56 Nevertheless, human and mouse trabecular meshwork share many common features and are more similar than those of rabbits and primates.

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FIGURE 1.9 Trabecular meshwork. A. This view of the posterior trabecular meshwork (delimited by arrows) contains six beams. Schlemm’s canal (SC) is lined by attenuated endothelial cells (arrowhead). Details of the boxed area are shown in C and E. Original magnification × 8100. B. The anterior trabecular meshwork (delimited by arrows) contains only two to three beams (compare to A). Typical giant vacuoles (V) occur on the anterior chamber side of Schlemm’s canal. Original magnification × 15,000. C. Enlarged view of boxed area in A shows details of trabecular beam structure. The thin flattened endothelial cells (arrow) cover a collagenous (C) and elastic tissue (arrowheads) core. Original magnification × 15,000. D. Midway between the anterior and posterior trabecular meshwork (arrows) there are numerous trabecular beams and giant vacuoles (V). Erythrocytes (E) are often observed in Schlemm’s canal (SC). Original magnification × 8100. E. Higher magnification of C showing details of the collagen core of the beams (*) and the elastic tissue (arrows). Original magnification × 30,000. F. The smooth muscle (M) of the ciliary body begins near the posterior termination of Schlemm’s canal. The muscle cells are filled with actin filaments and contain large collections of pinocytotic vesicles (arrow) at the cell surface. Original magnification × 15,000. (Figures D and E, courtesy of Smith, R. et al., BioMed Central, 1:3–14, 2001.)

LENS In comparison to the primate eye, the mouse lens is proportionately quite large, rounded, and occupies about 75% of the intraocular space (Figure 1.10).57,58 The lens is enclosed by the lens capsule, a homogeneous acellular structure that resembles basal lamina. In mice, the anterior lens capsule is about twice the thickness of the posterior capsule. Ultrastructural studies demonstrate a lamellar structure at high magnification.45,59,60 The presence of a zonular lamella in the superficial lens capsule has been suggested, but it appears to exist only near the lens equator and probably represents the gradual merging of the zonular fibers with the capsule.60-63 The zonules and superficial lens capsule contain abundant fibrillin, a high-molecular-weight glycoprotein.64 The lens epithelium lies directly beneath the lens capsule and contains numerous small mitochondria, a Golgi apparatus, and small amounts of endoplasmic reticulum (see Figure 1.10).45,65,66 A filamentous material of unknown composition is abundant in the basilar cytoplasm of lens epithelial cells.65 Anteriorly, the lens epithelium forms a monolayer of cuboidal cells with prominent lateral interdigitations. At the equator of the lens the epithelium becomes more columnar. Because the equator is the germinative zone for lens fibers, mitotic activity is most prominent in this location. Under normal circumstances, there are no lens epithelial cells beneath the lens capsule posterior to the germinative zone. Desmosomes and gap junctions connect the lateral cell membranes of lens epithelium in some animals.45,60 In mice, the only specialized lens epithelial cell contacts described are gap junctions.65 The epithelial cells of the equatorial germinative zone produce the lens fibers that make up the lens cortex. In the mouse, it is estimated that there are about 5000 stem cells at the lens equator that divide every 17 to 20 days and are responsible for lens fiber production.66 Unlike other epithelial structures, all cells produced by the lens epithelium persist throughout life. An epithelial cell differentiating to form a lens fiber begins by elongating anteriorly and posteriorly. Very quickly, the bulk of the cytoplasmic structures are eliminated, including mitochondria, endoplasmic reticulum, Golgi, and the nucleus.65 At the same time this is happening, extensive elaboration of the plasma membrane and cytoskeleton occurs. There is also rapid synthesis of specialized lens crystallins (lens proteins).67 The cytoplasm that remains is finely granular, and lacks structural features.60 The interface between mature lens fibers is characterized by several types of gap junctions and a complex array of “plug and socket” physical interconnections that can only be appreciated with scanning and freeze-fracture electron microscopy.60,68 The lens fibers vary in length, but may extend from the anterior to posterior poles of the lens. In a human eye, individual lens fibers may be as long as 7 mm; comparable figures for mice are unavailable. There is a tendency for the anterior and posterior cellular apices to come together to form a complex suture line.45,60

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FIGURE 1.10 Lens. A and B. The anterior lens capsule (arrow) is thick and covers the lens epithelium (arrowheads). The superficial lens cortex is indicated by C. Original magnification × 630. C. At the lens equator, the lens epithelial nuclei form a bow that curves into the cortex. Immediately posterior to the equator, the lens capsule (arrowhead) becomes thinner. Original magnification × 400. D. There is no lens epithelium beneath the thin posterior lens capsule (arrow). The cortex is indicated by C. Original magnification × 630. E. The anterior lens capsule (C) at this magnification is featureless, although fine fibrils can be identified at high magnification. The lens epithelium (E), directly beneath the capsule, has extensive cell membrane folding between adjacent cells. The lens cortex (CO) is acellular. F. Posterior to the lens equator, the capsule (C) is thinner and lens epithelial cells are absent. The lens cortical fibers develop elaborate cell membrane folding (arrows) that form “plug-and-socket” attachment points between adjacent fibers. E and F original magnification × 12,000.

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The zonules hold the lens in position and arise from the surface of the nonpigmented ciliary epithelium where they attach to the basal lamina of the cell. The zonules are made of bands of fibrils 10 to 12 nm in diameter with a periodicity of 11 to 18 nm in human eyes. The fibers merge with the superficial layer of the lens capsule, where they can be followed for only a short distance.60-63 The exact composition of the zonules remains uncertain, although fibrillin is an important component of both zonules and lens capsule.69

REFERENCES 1. Sundberg, J.P., Ed., Handbook of Mouse Mutations with Skin and Hair Abnormalities, CRC Press, Boca Raton, FL, 1994. 2. Sundberg, J.P., Hogan M.E., and King L.E., Normal biology and ageing changes of skin and hair, in Mohr, U. et al., Eds., Pathobiology of the Aging Mouse, ILSI Press, Washington, D.C., 1996, 303. 3. Michael, M.I. et al., Normal development of the eye-lids in the mouse, Folia Morph. (Praha), 36:53, 1988. 4. Yamashita, T., The spatial aspect and fine structure of the tarsal muscle of the mouse, J. Juzen Med. Soc., 88:1, 1979. 5. Yamashita, T., Takahashi, A., and Honjin, R., The spatial aspect and fine structure of the orbital muscle of the mouse, Okajimus Folia Anat. Jpn., 56:383, 1980. 6. Yamashita, T. and Honjin, R., Fine structure, origin, and distribution density of the autonomic nerve endings in the tarsal muscle in the eyelid of the mouse, Cell Tissue Res., 222:459, 1982. 7. Cook, M.J., The Anatomy of the Laboratory Mouse, Academic Press, London, 1965. 8. Popescu, P., Rajtova, V., and Horak, J., A Colour Atlas of the Anatomy of Small Laboratory Animals, Wolfe Publishing, London, 1992. 9. Sundberg, J.P. et al., Suppurative conjunctivitis and ulcerative blepharitis in 129/J mice, Lab. Animal Sci., 41:516, 1991. 10. Smith, R.S., Montagutelli, X., and Sundberg, J.P., Ulcerative blepharitis in aging inbred mice, in Mohr, U. et al., Eds., Pathobiology of the Aging Mouse, ILSI Press, Washington, D.C., 1996, 31. 11. Michael, M.I. et al., Normal postnatal development of the mouse eye, Folia Morphol., 36:125, 1988. 12. Wei, Z. et al., Label-retaining cells are preferentially located in fornical epithelium: implications on conjunctival epithelial homeostasis, Invest. Ophthalmol. Vis. Sci., 36:236, 1995. 13. Wirtschafter, J.D. et al., Mucocutaneous junction as the major source of replacement conjunctival epithelial cells, Invest. Ophthalmol. Vis. Sci., 40:3138, 1999. 14. Smith, R.S., The development of mast cells in the vascularized cornea, Arch. Ophthalmol., 66:383, 1961. 15. Nichols, B.A., Conjunctiva, Microsc. Res. Tech., 33:296, 1996. 16. Sundberg, J.P. et al., Myoepitheliomas in inbred laboratory mice, Vet. Pathol., 28:313, 1991. 17. Green, E.L., Ed., Biology of the Laboratory Mouse, Dover Publications, New York, 1966. 18. Tiffany, J.M., The role of Meibomian secretion in tears, Trans. Ophthalmol. Soc. U.K., 104:396, 1985. 19. Bron, A.J. and Tiffany, J.M., The Meibomian glands and tear film lipids, Adv. Exp. Med. Biol., 438:349, 1998. 20. Nakano, T., Postnatal transformation of the epithelium lining the nasolacrimal duct of the mouse, Z. Mikrosk. Anat. Forsch., 104:666, 1990. 21. Hay, E.D., Development of the vertebrate cornea, Int. Rev. Cytol., 63:263, 1980. 22. Hazlett, L.D. et al., Desquamation of the corneal epithelium in the immature mouse: a scanning and transmission microscopy study, Exp. Eye Res., 31:21, 1980. 23. Beuerman, R.W. and Pedoza, L., Ultrastructure of the human cornea, Microsc. Res. Tech., 33:320, 1996. 24. Oh, S.P., Griffith, C.M., and Olsen, B.R., Tissue-specific expression of Type XII collagen during mouse embryonic development, Dev. Dyn., 196:37, 1993. 25. Hazlett, L.D., Corneal and ocular surface histochemistry, Prog. Histochem. Cytochem., 25:1, 1993. 26. Rehbinder, C., Fine structure of the mouse cornea, Z. Versuchstierk., 20:28, 1978.

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Systematic Evaluation of the Mouse Eye: Anatomy, Pathology, and Biomethods 27. Haustein, J., On the ultrastructure of the developing and adult mouse corneal stroma, Anat. Embryol., 168:291, 1983. 28. LaVail, J.H., Johnson, W.E., and Spencer, L.C., Immunohistochemical identification of trigeminal neurons that innervate the mouse cornea: relevance to intercellular spread of herpes simplex virus, J. Comp. Neurol., 327:133, 1993. 29. Smith, R.S. et al., Haploinsufficiency of the transcription factors FOXC1 and FOXC2 results in aberrant ocular development, Hum. Mol. Genet., 9:1021, 2000. 30. Polack, F.M., Smelser, G.K., and Rose, J., Long-term survival of isotopically labelled stromal and endothelial cells in corneal homografts, Am. J. Ophthalmol., 57:67, 1964. 31. Schermer, A., Galvin, S., and Sun, T.T., Differentiation-related expression of a major 64K corneal keratin in vivo and in culture suggest limbal location for corneal epithelial stem cells, J. Cell Biol., 103:49, 1986. 32. Cotsarelis, G., Sun, T.T., and Lavker, R.M., Label-retaining cells reside in the bulge area of pilosebaceous unit: implications for follicular stem cells, hair cycle, and skin carcinogenesis, Cell, 61:1329, 1990. 33. Klareskog, L. et al., Expression of Ia-like antigen molecules on cells in the corneal epithelium, Invest. Ophthalmol. Vis. Sci., 18:10, 1979. 34. Sundberg, J.P., Boggess, D., and Sundberg, B.A., Epidermal dendritic cell populations in the flaky skin mutant mouse, Immunol. Invest., 22:389, 1993. 35. Rodrigues, M.M. et al., Langerhans cells in the normal conjunctiva and peripheral cornea of selected species, Invest. Ophthalmol. Vis. Sci., 21:759, 1981. 36. Rowden, G., Expression of Ia antigens on Langerhans cells in mice, guinea pigs, and man, J. Invest. Dermatol., 175:22, 980. 37. Streilein, J.W., Toews, G.B., and Bergstresser, P.R., Langerhans cells: functional aspects revealed by in vivo grafting studies, J. Invest. Dermatol., 75:17, 1980. 38. McLeish, W. et al., Immunobiology of Langerhans cells on the ocular surface. II. Role of central corneal Langerhans cells in stromal keratitis following experimental HSV-1 infection in mice, Reg. Immunol., 2:236, 1989. 39. Morrison, J.C. et al., Limbal microvasculature of the rat eye, Invest. Ophthalmol. Vis. Sci., 36:751, 1995. 40. Beamer, W.G. et al., Genetic variability in adult bone density among inbred strains of mice, Bone, 18:397, 1996. 41. Freddo, T.F., Ultrastructure of the iris, Microsc. Res. Tech., 33:369, 1996. 42. Smith, R.S., Ultrastructural studies of the blood-aqueous barrier. I. Transport of an electron-dense tracer in the iris and ciliary body of the mouse, Am. J. Ophthalmol., 71:1066, 1971. 43. Smith, R.S. and Rudt, L.A., Ultrastructural studies of the blood-aqueous barrier, Am. J. Ophthalmol., 76:937, 1973. 44. McMenamin, P.G. et al., Immunomorphologic studies of macrophages and MHC class II-positive dendritic cells in the iris and ciliary body of the rat, mouse, and human eye, Invest. Ophthalmol. Vis. Sci., 35:3234, 1994. 45. Hogan, M.J., Alvarado, J.A., and Weddell, J.E., Histology of the Human Eye, W.B. Saunders, Philadelphia, 1971. 46. Zimmerman, L.E. and Fine, B.S., Production of hyaluronic acid by cysts and tumors of the ciliary body, Arch. Ophthalmol., 72:365, 1964. 47. Haddad, A. et al., The origin of the intrinsic glycoproteins of the rabbit vitreous body: an immunohistochemical and autoradiographic study, Exp. Eye Res., 50:555, 1990. 48. Filho, R.B., Laicine, E.M., and Haddad, A., Biochemical studies on the secretion of glycoproteins by isolated ciliary body of rabbits, Acta Ophthalmol. Scand., 74:343, 1996. 49. Tripathi, B.J. and Tripathi, R.C., Neural crest origin of human trabecular meshwork and its implications for the pathogenesis of glaucoma, Am. J. Ophthalmol., 107:583, 1989. 50. Trainor, P.A. and Tam, P., Cranial paraxial mesoderm and neural crest cells of the mouse embryo: codistribution in the craniofacial mesenchyme but distinct segregation in branchial arches, Development, 121:2569, 1995. 51. Smith, R.S. et al., The mouse anterior chamber angle and trabecular meshwork develop without cell death, BMC Dev. Biol., 1:3, 2001.

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52. Tripathi, R.C., Mechanism of the aqueous outflow across the trabecular wall of Schlemm’s canal, Exp. Eye Res., 11:116, 1971. 53. Tripathi, R.C. and Tripathi, B.J., The mechanism of aqueous outflow in lower animals, Exp. Eye Res., 14:73, 1972. 54. Bill, A., Conventional and uveoscleral drainage of aqueous humor in the Cynomolgous monkey (Macaca irus) at normal and high intraocular pressures, Exp. Eye Res., 5:45, 1966. 55. Bill, A., A reply to R. Tripathi: uveoscleral drainage of aqueous humor, Exp. Eye Res., 25(Suppl.):309, 1977. 56. Gong, H., Tripathi, R.C., and Tripathi, B.J., Morphology of the aqueous outflow pathway, Microsc. Res. Tech., 33:336, 1996. 57. Zinn, K.M. and Mockel-Pohl, S., The lens and zonules, Int. Ophthalmol. Clin., 13:143, 1973. 58. Marshall, J., Beaconsfield, M., and Rothery, S., The anatomy and development of the human lens and zonules, Trans. Ophthalmol. Soc. U.K., 102:723, 1982. 59. Cohen, A.I., Electron microscopic observations on the lens of the neonatal albino mouse, Am. J. Anat., 103:219, 1958. 60. Kuszak, J.R., Peterson, K.L., and Brown, H.G., Electron microscopic observations of the crystalline lens, Microsc. Res. Tech., 33:441, 1996. 61. Farnsworth, P.N. et al., Surface ultrastructure of the human lens capsule and zonular attachments, Invest. Ophthalmol. Vis. Sci., 15:36, 1976. 62. Streeten, B.W., The zonular insertion: a scanning electron microscopic study, Invest. Ophthalmol. Vis. Sci., 16:364, 1977. 63. Streeten, B.W. et al., Immunohistochemical comparison of ocular zonules and microfibrils of elastic tissue, Invest. Ophthalmol. Vis. Sci., 21:130, 1981. 64. White, M.P. et al., Retinal degeneration in the nervous mutant mouse. II. Electron microscopic analysis, J. Comp. Neurol., 333:182, 1993. 65. Rafferty, N.S. and Esson, E.A., An electron-microscopic study of adult mouse lens: some ultrastructural specializations, J. Ultrastruct. Res., 46:239, 1974. 66. Rafferty, N.S. and Rafferty, K.A., Cell population kinetics of the mouse lens epithelium, J. Cell. Physiol., 107:309, 1981. 67. Wistow, G.J. and Piatigorsky, J., Len crystallins: the evolution and expression of proteins for a highly specialized tissue, Annu. Rev. Biochem., 57:479, 1988. 68. Lo, W. and Reese, T.S., Multiple structural types of gap junctions in mouse lens, J. Cell Sci., 106:227, 1993. 69. Wheatley, H.M. et al., Immunohistochemical localization of fibrillin in human ocular tissues, Arch. Ophthalmol., 113:103, 1995.

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Posterior Segment and Orbit Richard S. Smith, Simon W. M. John, and Patsy M. Nishina

CONTENTS Vitreous . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .25 Posterior Uvea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .27 Choroid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .27 Retina . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .27 Pars Plana . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .27 Retinal Organization and Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .27 Retinal Pigment Epithelium and Bruch’s Membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .29 Photoreceptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .31 Outer Plexiform Layer and Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .31 Inner Nuclear Layer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .33 Inner Plexiform Layer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .35 Ganglion Cell Layer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .35 Nerve Fiber Layer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .35 Retinal Vascular System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .36 Optic Nerve and Meninges . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .36 Orbit and Extraocular Muscles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .38 Orbit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .38 Extraocular Muscles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .38 Orbital Vascular Supply . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .39 Harderian Gland . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .40 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .41

VITREOUS The vitreous occupies the posterior portion of the eye between the lens and retina. This compartment is small in mouse eyes relative to primates. With light microscopy and routine stains, the vitreous chamber appears empty, because its content is mostly water. However, special stains for glycosaminoglycans and hyaluronic acid demonstrate the presence of these complex substances in the vitreous compartment. The highest concentrations are found near the retinal surface and in the vitreous directly posterior to the lens.1-6 Light microscopy hints that there is an intimate attachment between the peripheral vitreous and retina. Transmission electron microscopy reveals the presence of irregularly arranged collagen fibers that are concentrated in the peripheral vitreous (Figure 2.1). The vitreous collagen fibrils are attached to the basal lamina of the retina and ciliary body.1-6 Attachments of vitreous to the posterior lens capsule also occur in human eyes.7 However, the ease with which the mouse lens can be physically removed suggests that vitreolenticular attachments are minimal or absent in mice. Reference is often made to the “anterior hyaloid membrane” of the vitreous in mouse1-3 and human eyes.7 The structure referred to is a focal condensation of collagen fibers at the anterior vitreous surface, rather than a true membrane.7 0-8493-0864-X/02/$0.00+$1.50 © 2002 by CRC Press LLC

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FIGURE 2.1 Vitreous and retina. A 2-week-old C57BL/6J mouse. Hyaloid vessels (V) still persist at this age. A hyalocyte (H) is located close to the retinal (R) surface. The hyalocyte has scanty cytoplasm and few cytoplasmic organelles. Vitreous collagen fibrils (arrowheads) are concentrated close to the retinal surface and are intimately related to the internal limiting membrane (arrow). Original magnification × 24,000.

Vitreous cells known as hyalocytes have been described in many animals,7 but have not previously been reported in the mouse. Hyalocytes are round cells with prominent nuclei that contain variable numbers of cytoplasmic granules thought to be secretory.7 Hyalocytes are more easily identified in younger mammals. A typical mouse hyalocyte is demonstrated in Figure 2.1. It has been suggested that hyalocytes belong to the monocyte/macrophage lineage because they express common antigens

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that include PTPRC (formerly CD45), ITGAL (formerly CD11a), and FCGR1 (formerly CD64). They differ from tissue macrophages in that they express S100 protein and do not express CD68.8 This suggests that hyalocytes are true vitreal cells and not casual visitors from the systemic circulation. Hyalocytes express messenger RNA for transforming growth factor-β (TGF-β) and hyalocyteconditioned media inhibits mitosis in retinal pigment epithelial cells. It was suggested, on this basis, that one hyalocyte role might be control of cell proliferation in intraocular disease.9

POSTERIOR UVEA CHOROID The anterior uvea (Chapter 1) includes the iris and ciliary body; the choroid is the posterior uvea. The choroid (Figure 2.2) is a highly vascular tissue that provides the blood supply for the external layers of the retina (retinal pigment epithelium through the outer portion of the inner nuclear layer). In humans, the choroid receives its blood supply from the anterior and posterior ciliary arteries that are branches of the ophthalmic artery. Branches of the ciliary arteries are found more externally in the choroid, as are the venules of the venous drainage system. The veins leave the eye via three to four vortex veins, which enter the orbital venous sinus. It is likely that a similar pattern exists in mice, but that has not been confirmed. The choriocapillaris is the innermost vascular layer and here the endothelial cells are thin and heavily fenestrated, similar to the ciliary body. The basal lamina of the choriocapillaris also forms the most external part of Bruch’s membrane (see below). The nerve supply of the choroid comes from the short posterior ciliary nerves. In human eyes, ganglion cells have been identified in the choroidal stroma, suggesting an intraocular autonomic supply.7 Dendritic melanocytes and fibroblasts are abundant in the choroidal stroma, which is quite thin in mice. There are numerous mast cells in the choroid that can be identified by stains such as acidic toluidine blue.10 Macrophages and lymphocytes have also been identified in the choroid.7

RETINA PARS PLANA The junction between the retina and ciliary body is quite lengthy in primates.7 Here, the retina thins rapidly from the multiple layers described below to two layers of cells, the external layer pigmented and the internal nonpigmented. This scalloped border of retinal termination is known as the ora serrata. The bilayer of cells continues over the posterior aspect of the ciliary body where it is known as the pars plana ciliaris. Numerous ciliary processes (see Figures 1.8 and 1.9, Chapter 1) are formed (pars plicata), over the ciliary body. These may differ considerably in appearance as a result of the angle of section. The ora serrata region described in humans is similar in mice (see Figure 2.2), but the pars plana is much shorter, usually not more than 12 to 16 cells wide.

RETINAL ORGANIZATION AND FUNCTION The neural retina is the most complex substructure of the eye and, in many ways, functions anatomically and physiologically as an outpost of the central nervous system (Figures 2.2 through 2.4). The retina develops from two layers of neuroepithelium of the optic cup. The inner layer of the optic cup proliferates and undergoes a complex process of differentiation (Chapter 3) to form the neural retina. The outer layer of the optic cup becomes the retinal pigment epithelium and remains a monolayer.

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FIGURE 2.2 Choroid and retina. A. The interface between the vitreous and retina is the internal limiting membrane (arrowhead). There is a single layer of ganglion cells (G) in the equatorial retina and the nerve fiber layer is not easily identified (see Figure 2.3). The inner plexiform layer (IP) is nearly as wide as the outer nuclear layer (ON). The inner nuclear layer (IN) includes the bipolar, amacrine, horizontal, and Muller cells. The external limiting membrane (arrow) separates the outer nuclear layer from the photoreceptor inner segments (IS) and outer segments (OS). Original magnification × 630. B. The choroid (C) contains many dendritic melanocytes, as well as vascular channels; the choriocapillaris is adjacent to the retinal pigment epithelium (arrowhead), while arterioles and venules lie closer to the sclera (S). Original magnification × 400. C. The basal portion of a retinal pigment epithelial cell (*) has prominent cytoplasmic membrane folding. The choriocapillaris is located close to the RPE and its endothelial lining is fenestrated. The RPE and choriocapillaris are separated by Bruch’s membrane (arrow). Inset: The elastic tissue component of Bruch’s membrane (arrow) lies between the basal laminas of the RPE and the endothelium of the choriocapillaris. Basal infoldings of the RPE (*) are prominent. Original magnification × 6000, inset × 15,000. D. At the pars plana (arrows), the retina (R) thins abruptly to a single layer of nonpigmented epithelial cells that continue anteriorly as the nonpigmented ciliary epithelium of the ciliary body (CB). Original magnification × 630. E. The tips of the photoreceptor outer segments are phagocytosed by the RPE. In some cases, the folded lamellae of the outer segments are recognizable (arrowhead), but as digestion proceeds, the phagosomes become more difficult to recognize (arrows). Original magnification × 6000.

The visual cascade begins when light initiates complex chemical changes in the visual pigments, generating an electrical impulse that travels down the photoreceptor axon to synapse with cells of the inner nuclear layer. The photoreceptors act as the peripheral sense organ, similar to those of other senses. After the initial synapse, there is extensive lateral and vertical modification of the signal by the cells of the inner nuclear and ganglion cell layers. For example, the initial recognition of edge effects occurs in the retina, affecting interpretation of what reaches the visual cortex.7

RETINAL PIGMENT EPITHELIUM AND BRUCH’S MEMBRANE The retina is separated from the choroid by the monolayer of retinal pigment epithelium (RPE). In pigmented mouse strains, numerous melanosomes are present (see Figure 2.2). The individual melanosome is relatively ovoid in shape compared with those found in dendritic melanocytes in the choroid or iris. Unpigmented melanosomes (premelanosomes) are present in albino mice and are most easily seen by electron microscopy. Mitochondria are abundant in the basal aspect of the cell, which reflects the intense metabolic activity of the RPE. Membrane-enclosed lamellar inclusions, called phagosomes, are found in the cytoplasm of the RPE in all vertebrate eyes and represent fragments of photoreceptor outer segments that are being recycled for renewal of these structures (see Figure 2.2).11-14 Microvilli extend from the apical cell membrane of the RPE and surround the outer third of the photoreceptor outer segments. The interphotoreceptor matrix in most mammals consists of glycosaminoclycans (hyaluronan and others), glycoproteins, and glycoconjugates that fill the spaces between adjacent photoreceptors and RPE villi. Hyaluronan has been reported to be absent from the retina of some mouse strains.7,15 The lateral cell membranes have simple interdigitations and apical tight junctions that are impermeable to small-molecular-weight tracers.16 The basal cell membrane of the RPE is characterized by prominent infoldings that reflect the role of the cell in transport of substances between the choroidal circulation and the retina. The basal lamina of the RPE forms the innermost layer of Bruch’s membrane. The outermost layer of Bruch’s membrane is the basal lamina of the choriocapillaris. Between these two basal laminae, there are delicate layers of collagen and elastic tissue. The layering of Bruch’s membrane is present, but less obvious in mice than in primates.

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FIGURE 2.3 Retina and optic nerve. A. Near the optic nerve, the retinal ganglion cell layer (arrowhead) is two to three cells thick. The nerve fiber layer (arrow) is easily identified. Original magnification × 630. B. Posterior to the equator of the eye, the outer nuclear layer (ONL) is normally 10 to 12 cells thick. The photoreceptor inner segments (IS) are densely packed and stain a bit more intensely than the outer segments (OS). There is normally no space between the outer segments and the retinal pigment epithelium (*). Original magnification × 630. C. Close to the optic nerve, there is obvious thickening of the nerve fiber layer (arrow). Original magnification × 400. D. At the point where the nerve fiber layer enters the optic nerve (ON), all retinal layers abruptly terminate (arrows). Mice have a well-defined lamina cribrosa (arrowheads) in which collagen and cell nuclei are arranged perpendicular to the long axis of the optic nerve. Original magnification × 200. E. With most stains, the only identifiable parts of the Muller cell are the internal and external limiting membranes. When glial fibrillary acid protein immunohistochemical stains are used, both the internal limiting membrane (arrowhead) and the vertically arranged Muller cell cytoplasm (arrows) are revealed. The photoreceptor nuclei are absent, as this mouse has advanced retinal degeneration. Original magnification × 400. F. The laminar structure of the nerve fiber layer (NF) is evident as it enters the optic nerve (ON). The termination of the retina, retinal pigment epithelium, and choroid (arrow) is often tapered. Original magnification × 630.

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PHOTORECEPTORS The outer nuclear layer of mice is 10 to 12 nuclei thick (see Figures 2.2 through 2.4). There is very little perinuclear cytoplasm. Delicate Muller cell cytoplasmic processes extend vertically across the outer nuclear layer. Approximately 95% of the photoreceptor cells in the mouse retina are rods.17,18 Two types of cone cells have been identified in mouse retina, although they can only be distinguished by immunohistochemistry or electroretinography (see Chapter 14).17,19,20 This finding suggests that mice have at least dichromatic vision.17 It also supports the hypothesis that the same complex interaction between rod and cone photoreceptors that occurs in primates may also function in mice (see below).21-25 Generally, the nuclei of rod photoreceptors are densely basophilic with unevenly distributed nucleoplasm. Cone photoreceptors have slightly larger nuclei, stain less densely, and are most often located near the external limiting membrane. The photoreceptor nuclei are separated from the photoreceptor inner and outer segments by the external limiting membrane (see Figure 2.2). Those unfamiliar with the eye should not be confused by this firmly established but ill-conceived name. The external limiting membrane is not external, limits nothing, and is not a membrane. This nevertheless prominent anatomic feature is the result of the regularly arranged adherent junctional complexes between adjacent photoreceptors and Muller cells and is easily visualized by light microscopy.7,26 The photoreceptor inner and outer segments (see Figures 2.3 and 2.4) are located just external to the external limiting membrane. The broad inner segments are divided into an outer ellipsoid portion that is packed with mitochondria and an inner myoid portion that contains abundant glycogen and ribosomes. These features are consistent with the high level of metabolic and synthetic activity of the inner segments. The inner segments are connected to the narrower outer segments by a connecting cilium with nine pairs of tubules, easily visualized by electron microscopy (see Figure 2.4). This serves as a reminder that the outer segments are modified cilia, reflecting their origin from ciliated neuroepithelium. The connecting cilium in photoreceptors is more than a simple mechanical linkage between inner and outer segments. The constant renewal of the outer segment lamellae requires synthesis and transport of protein. Photoreceptor protein synthesis and its transport through the connecting cilium have been demonstrated in several species (rat, mouse, frog, and monkey).11-13,27-29 Directly external to the cilial structures, the plasma membrane of the photoreceptor forms numerous folds, producing stacks of photoreceptor lamellae surrounded by a plasma membrane. The inner and outer segments of rod photoreceptors have a uniform diameter, whereas the inner segments of cones have a tapered shape, broader on the cilial side. Adjacent photoreceptor outer segments are separated by fine processes that extend from the Muller cells.26

OUTER PLEXIFORM LAYER AND SYNAPSES The thin outer plexiform layer is the layer of synapses (see Figures 2.2 and 2.4) between the photoreceptors and the neural cells of the inner nuclear layer. The actual synapses are clearly distinguished only by electron microscopy. The rod and cone axons arise from the cell bodies of the respective photoreceptors and are filled with neurofilaments. The rod axon terminates in an ovoid rod spherule, which is the site of the synapse. The major portion of the rod spherule is presynaptic and the principal cytoplasmic feature is the presynaptic vesicles. The dendrites of bipolar and horizontal cells invaginate the plasma membrane of the rod spherule, forming the synaptic cleft. Usually, multiple dendritic processes form the synapse and they often originate from multiple cells in the inner nuclear layer. Postsynaptic vesicles and mitochondria are found in the dendritic cell processes of the rod spherule.7,30 The synaptic area may have multiple synaptic ribbons, depending on the number of nerve processes that reach the cell. In addition to photoreceptors, ribbon synapses are found in bipolar cells, the hair cells of the organ of Corti, the vestibular organ, and in the intrinsic cells of the pineal gland.31 The synaptic ribbon is a densely osmiophilic lamellar structure, always closely aligned with

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FIGURE 2.4. Retina. A. The outer segments (OS) of the photoreceptors are in direct contact with the RPE (PE). The inner segments (IS) connect the photoreceptor cell bodies of the outer nuclear layer to the outer segments. Original magnification × 6000. B. The inner segments contain many mitochondria (arrowhead). The outer nuclear layer (ONL) contains numerous rod nuclei (arrow) with densely staining chromatin. Cone nuclei (open arrow) tend to be larger with less nuclear chromatin. Original magnification × 6000. C. Synapses between the photoreceptors from the outer nuclear layer (ONL) and the cells of the inner nuclear layer (INL) are located in the inner plexiform layer (IPL). Inset: A connecting cilium (arrow) joins the inner to the outer segment. Original magnification × 8100, inset × 15,000. D. At higher magnification the complex synaptic connections (arrows) of the photoreceptors are located in the inner plexiform layer. A synaptic ribbon is indicated by the arrowhead. Original magnification × 12,000.

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the dense arciform density, a part of the synaptic ribbon complex. It has been suggested that synaptic ribbons act to convey the synaptic vesicles to the presynaptic membrane. An alternative hypothesis is that quantities of synaptic vesicles are linked together by a specific phosphoprotein that immobilizes them.31 Recently, it has been demonstrated that the morphological appearance of synaptic ribbons varies with the time of day and intensity of illumination.32 Clearly, much remains unknown concerning their function. The cone axon terminates in a broad, pyramidal structure known as the cone pedicle, which is broad at its base. Because of the small population of cones in mice (3 to 5% of total photoreceptors), these structures are much less common than in primates. The arrangement of synaptic vesicles and synaptic ribbons is similar to that described in the rods. Cone synapses consist of multiple, deeply invaginated structures known as triads. Both horizontal and bipolar cell processes connect with the cones. Horizontal cell processes extend laterally and synapse with photoreceptor and bipolar cells in the outer plexiform layer.21-24 Bipolar cells also makes surface contacts with the cone pedicle. Finally, there are interphotoreceptor contacts between both rods and cones. It should be emphasized that the functional organization is much more complex than first inspection might indicate.7,21-25,30,33

INNER NUCLEAR LAYER The inner nuclear layer (Figures 2.3 through 2.5) is a dense layer that is six to nine cells thick in the peripapillary retina and includes the nuclei of bipolar, horizontal, amacrine, and Muller cells. The bipolar cells that connect the photoreceptor and ganglion cell layers are most abundant. Several types of bipolar cell have been described, including midget, flat, and rod bipolar cells. The latter are the only bipolar cells that connect to the rods. Each rod bipolar cell contacts the rod spherule and also connects with from one to four ganglion cells. The midget bipolar cells connect with the triads of the cone pedicle and synapse with a single midget ganglion cell. They do not contact adjacent bipolar cells. The flat bipolar cells are polysynaptic cells that connect primarily with cones.7,30,33 This classification requires the use of special heavy metal stains as these cells are most easily identified by the nature of their synaptic connections.7 Studies using labeling by retrograde axonal transport and immunohistochemistry provide additional information concerning the complexity of cell-to-cell contacts in the retina.34 The incredible intricacy of interconnections is suggested by the data from a three-dimensional electron microscopic reconstruction. Rod terminals may have horizontal cell to horizontal cell, horizontal to bipolar cell, rod to bipolar cell, cone to rod, and cone to bipolar cell synapses and in various combinations the synaptic connections may be depolarizing or hyperpolarizing.23 A single reconstructed cone terminal received 60 terminal branches from 19 neurons with nearly 150 synapses.21 A detailed analysis of this complexity is beyond the scope of this discussion. The interested investigator is referred to the extensive review of the subject by Sjostrand.21-24 Horizontal and amacrine cells are also found in the inner nuclear layer. Horizontal cells are similar in appearance to bipolar cells, except for a cytoplasmic inclusion known as Kollmer’s crystalloid, the function of which is unknown. The inclusion stains densely basophilic and can be demonstrated by light microscopy, but has not been reported in mice. As previously noted, horizontal cells connect to cone pedicles and rod spherules in a complex pattern. Amacrine cells are larger than horizontal cells, have a highly indented nucleus with prominent nucleolus, and their nuclei stain less densely than those of bipolar cells. A number of subtypes of amacrine cells have been described.7,33 Both amacrine and horizontal cells are responsible for lateral integration of neurons, and each may connect to many photoreceptors or ganglion cells as well as synapse with each other and with adjacent bipolar cells. The processes of amacrine cells extend laterally, connecting to bipolar and ganglion cells in the inner plexiform layer.21 The nuclei of Muller cells (see Figures 2.2 through 2.5) are also located in the inner nuclear layer. The cytoplasm of Muller cells stains more darkly than other cells in the inner nuclear layer. A

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complex array of Muller cell cytoplasmic processes extends from the retinal surface to the villous processes that lie just beyond the external limiting membrane. Muller cells, which fill retinal space unoccupied by neurons, are a source of glucose and glycogen, and help to maintain retinal structural integrity.26 Most importantly, Muller cells are known to be responsible for removal of glutamate from the extracellular space. Because glutamate likely plays an important role in glaucoma, Muller cell

FIGURE 2.5 Inner retina. A. A retinal ganglion cell (G) is located close to the internal limiting membrane (arrow). A larger vascular channel (V) is filled with erythrocytes. B. Several classes of cells are located in the inner nuclear layer. Amacrine cells (A) are large with typically indented nuclei. Horizontal cells (H) also have large nuclei and abundant cytoplasm. Bipolar cells (B) have smaller nuclei, with a variable morphology. The cytoplasm of Muller cells (M) is scanty and long cytoplasmic processes (arrow) extend toward the inner and outer retina, terminating on the internal and external limiting membranes. Original magnification × 15,000.

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function may be critical for survival of other retinal neurons (reviewed in Carter-Dawson et al.35). The complex ultrastructural arrangement of the Muller cell cytoplasm in human and primate eyes is reviewed in Hogan et al.7 Limited observation suggests a similar complexity in mice (R. Smith, unpublished observations).

INNER PLEXIFORM LAYER Synapses between ganglion, bipolar, and amacrine cells occur in the inner plexiform layer. Muller cell processes and an occasional astrocyte are also found here.26 Although usually free of nuclei, a sporadic displaced cell from the ganglion cell or inner nuclear layer may be found in the inner plexiform layer. Neurotubules and neurofilaments are abundant in bipolar cell axons. Synaptic ribbons are characteristic of the bipolar cell synapses, which makes them easy to identify in electron microscopic preparations. Both ganglion cell dendrites and amacrine cell processes are found at the bipolar synapse. This arrangement is known as a dyad.33 Amacrine cell processes are identified by cytoplasm that is less dense than that of bipolar cell axons. Axosomatic and axodendritic contacts occur between bipolar, ganglion, and amacrine cells, forming another complex array of intercellular connections.7,30,33,34

GANGLION CELL LAYER The retinal ganglion cells are characterized by large vesicular nuclei with prominent nucleoli and abundant cytoplasm. As with other neurons, the endoplasmic reticulum of ganglion cells tends to aggregate in large clumps that form the Nissl substance visible by light microscopy. Special silver staining techniques can be used to identify several different types of ganglion cells with specific synaptic connections.7 These cells cannot be distinguished by routine light or electron microscopy.7 Smaller oval nuclei with dense nuclear staining found in the ganglion cell layer most likely represent endothelial or pericyte nuclei from vascular channels. Amacrine cells have also been identified in the ganglion cell layer by immunohistochemistry, but cannot be definitively identified in routine sections. In a normal mouse retina, the ganglion cell nuclei are closely spaced and in the peripapillary area may be two to three cells thick. Ganglion cell density is greatest just temporal to the optic nerve (>8000/mm2) and least (20 cM. The method by which the interval is narrowed will depend greatly upon the contribution that a particular locus has on the observed phenotype. If a major QTL is identified that is responsible for the majority of the phenotypic variation within a particular cross, then the region can be narrowed by progeny testing of mice that are recombinant within the QTL/modifier interval. In the case of localizing a recessive QTL/modifier locus, F2 intercross or backcross progeny are tested for markers flanking the QTL/modifier. Each recombinant mouse is mated to a mouse that is homozygous for the QTL/modifier locus. If the recombinant chromosome contains the locus, then half the offspring will show the QTL phenotype. Typically, at least 20 offspring are phenotyped and the distribution should not be significantly different from the expected 1:1 ratio of affected to unaffected. The number of meioses tested will determine the size of the genetic interval. For example, if 1000 F2 mice are tested, the region containing the QTL could be theoretically as small as 0.05 cM in size. On the other hand, if multiple QTL/modifier loci are identified that have modest effects, interval specific congenic strains may need to be constructed to isolate the various loci. A more detailed discussion of the different methods of recombinant progeny testing, including the following congenic approach, is given by Darvasi.40 A congenic is a variant strain that is constructed by repeated backcrossing of a donor strain carrying a genomic region of interest to a recipient inbred strain for ten backcross generations with subsequent intercrossing.41 With each subsequent backcross generation, the percentage of donor genome declines by one half in unlinked regions, such that by ten backcross generations, 99.9% of the unlinked genomic regions are of recipient strain origin. The purpose of the congenic is to isolate the QTL/modifier so that any phenotypic variation that is observed between the congenic and the recipient parental strain must be a consequence of the gene encompassed by the congenic interval. The congenic can then be used in traditional recombinant crosses to narrow the genetic interval containing the QTL/modifier.

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Previously, congenics were constructed by selection for a particular phenotype. Donor mice exhibiting the phenotype desired were used as donor parentals at each backcross generation. Currently, congenics are constructed using marker-assisted selection. There are two extremes of this latter approach; one is simply to select donors for subsequent backcross generations that are heterozygous at markers flanking the QTL/modifier. The other approach is called speed congenic construction in which selection against unlinked donor genome is employed in addition to the selection for the locus of interest.42 As the name implies, rather than the normal ten backcross generations, the congenic strain can be constructed in fewer than five generations. This method, which is labor-intensive, requires genome-wide typing of a large number of progeny from the first backcross to allow one to select the donor mouse with the highest percentage of recipient genome background to use in the second backcross. Beyond the second backcross generation, marker analysis is used only for selection of the region of interest, because counter selection is no longer efficient. With either of the two approaches, it is prudent to check during the construction process whether the phenotype in question is still expressed in the incipient congenic. Pitfalls may occur during congenic construction, but there are potential methods of circumventing these problems.31 These problems include but are not limited to loss of the phenotypic expression of the QTL/modifier due to unaccounted for epistatic interactions, or due to a low contribution of the QTL to the phenotype in question. Once the region is reasonably narrowed and a physical resource has been identified (see discussion below), another genetic technique can be used to refine further the QTL/modifier location. Transgenic whole large insert clone (e.g., BACs, PACs, P1s) rescue43-45 has become a widely used approach that may potentially narrow the critical region to one large insert clone, a feat that is usually not achieved by recombinational mapping alone. The drawbacks are that the gene of interest must be entirely encompassed within one clone, special skills are needed to perform the pronuclear injections, and a cost is associated with these experiments. To obviate the potential of missing a gene at the end of a clone, clones with significant overlap are injected singly into the pronucleus of blastocysts. Alternatively, investigators have injected two overlapping BACs and have shown that homologous recombination of the BACs occurs upon transgene insertion.44

GENE IDENTIFICATION POSITIONAL CANDIDATES Numerous gene identification projects that began as a positional cloning effort have, after further refinement of a map position, been concluded by identifying a candidate responsible for the disease phenotype.46,47 If genes that are reasonable biological candidates map to the QTL/modifier region, oligonucleotide primer sets can be created that flank a portion of the gene (genomic DNA) likely to contain a sequence variation (i.e., introns). The primers can be used in a PCR assay to test if the candidate gene maps to the critical region. Either DNA from mice carrying recombinations within the critical region can be tested or the candidate gene can be mapped with respect to markers on the T31 mouse radiation hybrid panel (Research Genetics, AL). Alternatively, if a physical resource spanning the critical region is available (see below), the presence of the candidate gene in the large insert clones can be determined by PCR analysis. Finally, as genomic sequences of the human and mouse genome become available, computer databases can be searched to determine if a candidate gene is present within the critical region.48 If a gene of interest maps within the region that contains the QTL/modifier, mutation detection screening can ensue.

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POSITIONAL CLONING Currently, once positional candidates are exhausted, genes can be identified by the process of positional cloning, that is, identification of a gene based on positional information alone. The process involves refining the genetic interval (discussed above), assembling a physical contig with large insert clones across the critical interval, identification of transcripts within the assembled physical resource, mutation detection screening, and verification. Within the next few years as assembled and annotated sequences of the human and mouse genome become available, some of the steps that are currently necessary for the positional cloning process will become obsolete. For example, in most cases it will no longer be necessary to assemble a physical contig or identify transcripts using techniques such as direct cDNA selection49 or exon trapping.50 Therefore, these methodologies will not be discussed here.

ASSEMBLY, ANNOTATION/IDENTIFYING TRANSCRIPTS The availability of the sequences from the Human Genome Project has greatly facilitated positional cloning efforts. When a human sequence contig is available, transcripts can be located by searching for sequence similarities between the genomic sequence and entries in Expressed Sequence Tag (EST) databases. EST databases are collections of short sequences (tags, ~200 to 500 bps) obtained by randomly sequencing the ends of cDNA clones of libraries derived from many different tissues. Currently, there are ~3.6 million human and ~2 million mouse EST sequences in the databases. Most genes are represented by several EST entries, and often the complete cDNA sequence can be assembled from EST entries and such assemblies have been compiled in so-called gene indices like UNIGENE51 and TIGR Gene Index (http://www.tigr.org/tdb/mgi/). Since ESTs are derived from cDNA, these sequence tags represent the expressed sequences (transcripts) in these tissues. A fast sequence alignment algorithm, BLAST (http://www.ncbi.nlm.nih.gov/blast/), is used to search for sequences in the database that are similar to a query sequence.52 In addition, a combination of software programs designed to identify coding regions within genomic sequences can be used to predict genes. Burset and Guigo53 have shown that such a strategy maximizes the chance of identifying all transcripts in a given region. Several Web tools are available that carry out automated BLAST searches and gene predictions on submitted genomic sequences (annotation pipelines). One example is the NIX annotation pipeline (http://www.hgmp.mrc.ac.uk/Registered/Webapp/nix/). The output of this pipeline is a list (or graphical representation) of the potential coding sequences in the genomic fragment. Additional BLAST searches are then used to determine if those potential coding sequences that were identified can be expanded to, ideally, a full-length cDNA sequence (gene indices) or at least a significant portion of the cDNA sequence. Once a human cDNA is assembled, mouse EST databases can be searched to obtain the mouse homologues. In cases where the mouse EST sequences cannot be extended by in silica methods, full (or nearly full) length sequences must be obtained experimentally through screening of cDNA libraries or 5′ and 3′ RACE reactions (Rapid Amplification of cDNA Ends).54

MUTATION DETECTION To begin searching for mutations, it is useful to carry out Northern blot analysis to determine if changes in the level of expression of a transcript or alterations in the transcript size are observed. Apart from being due to promoter or splice site mutations, such changes can also hint at larger-scale genomic alterations such as retroviral insertions or deletions. Southern analysis can be used to detect these in genomic DNA through changes in the restriction digest pattern between mutant (the congenic carrying the QTL) and wild-type (the background strain in which the congenic was constructed). After obtaining full-length cDNA sequences, oligonucleotide primers can be constructed that allow amplification of the whole coding region by reverse transcriptase PCR (RT-PCR) from mRNA obtained from the congenic and the background strain. Sequence comparison will identify coding

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region mutations such as missense (amino acids changes) and nonsense (premature stop codons) mutations. Splice site mutations and deletions are recognized by missing coding sequences. If a sequence change is found, it should be confirmed at the genomic level by constructing oligonucleotide primers that allow amplification of the corresponding genomic area. If a change in transcript level was observed, but no coding sequence differences were detected, then regulatory sequences may be altered and the regions 5′ to the gene, introns, and 3′ noncoding regions will need to be examined.

VERIFICATION Particularly in the identification of QTL, it is of prime importance to verify that the detected sequence change is indeed causing the observed phenotype. This is because it is commonly thought that QTL are due to sequence polymorphisms that are present in many inbred strains. An approach applicable to the verification of a recessively acting QTL is a transgenic rescue experiment. In this case the gene that carries the sequence polymorphism thought to cause the phenotype is expressed in its wild-type form, preferably under its own promoter, in the congenic carrying the QTL. If the transgene restores the wild-type phenotype to the congenic, we conclude that the correct gene underlying the QTL has been identified. In the case of a dominant acting QTL, one would make a transgenic animal carrying the altered form of the gene in the background strain to see if transgene expression can mimic the phenotype of the congenic strain. Another possibility is the creation of a null allele of the QTL (by inactivating the gene through the introduction of a deletion by homologous recombination) to induce, possibly, an even more severe phenotype. Alternatively, one can carry out a gene replacement experiment in which the polymorphism is introduced into the wild type gene by means of homologous recombination.

SUMMARY The mapping and subsequent identification of single gene mutations and, more recently, the analysis of targeted mutations were instrumental in improving our understanding of the cellular processes that are necessary for normal functioning of the eye and other organs. Although not all Mendelian eye diseases are identified and more work is still necessary, the time has come to begin to understand complex diseases and inheritance patterns. QTL analysis is highly effective in localizing genes that underlie complex traits. Although computer-driven approaches to finding QTL are beginning to be developed, the experimental approach will remain useful for some time. The completion of human genome sequencing, the progress on the mouse sequence, and the development of large-scale expression profiling methods hold great promise for translating QTL positions into knowledge about the genes underlying the trait.

REFERENCES 1. RetNet, Retinal Information Network. Laboratory for the Molecular Diagnosis of Inherited Eye Diseases, The University of Texas-Houston Health Science Center, 2001, available at http://www.sph.uth.tmc.edu/RetNet/. 2. OMIM, Online Mendelian Inheritance in Man. McKusick-Nathans Institute for Genetic Medicine, Johns Hopkins University, Baltimore, MD, and National Center for Biotechnology Information, National Library of Medicine, Bethesda, MD, 2001, available at http://www.ncbi.nlm.nih.gov/omim/. 3. Weleber, R.G. et al., Phenotypic variation including retinitis pigmentosa, pattern dystrophy, and fundus flavimaculatus in a single family with a deletion of codon 153 or 154 of the peripherin/RDS gene, Arch. Ophthalmol., 111:1531, 1993. 4. Kajiwara, K., Berson, E.L., and Dryja, T.P., Digenic retinitis pigmentosa due to mutations at the unlinked peripherin/RDS and ROM1 loci, Science, 264:1604, 1994.

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Systematic Evaluation of the Mouse Eye: Anatomy, Pathology, and Biomethods 5. Gibson, F. et al., A type VII myosin encoded by the mouse deafness gene shaker-1, Nature, 374:62, 1995. 6. Travis, G.H. et al., Identification of a photoreceptor-specific mRNA encoded by the gene responsible for retinal degeneration slow (rds), Nature, 338:70, 1989. 7. Pittler, S.J. and Baehr, W., Identification of a nonsense mutation in the rod photoreceptor cGMP phosphodiesterase beta-subunit gene of the rd mouse, Proc. Natl. Acad. Sci. U.S.A., 188:8322, 991. 8. Noben-Trauth, K. et al., A candidate gene for the mouse mutation tubby, Nature, 380:534, 1996. 9. Weil, D. et al., Defective myosin VIIA gene responsible for Usher syndrome type 1B, Nature, 374:60, 1995. 10. Kajiwara, K. et al., Mutations in the human retinal degeneration slow gene in autosomal dominant retinitis pigmentosa, Nature, 354:480, 1991. 11. McLaughlin, M.E. et al., Recessive mutations in the gene encoding the beta-subunit of rod phosphodiesterase in patients with retinitis pigmentosa, Nat. Genet., 4:130, 1993. 12. Banerjee, P. et al., TULP1 mutation in two extended Dominican kindreds with autosomal recessive retinitis pigmentosa, Nat. Genet., 18:177, 1998. 13. Fitzsimons, R.B., Gurwin, E.B., and Bird, A.C., Retinal vascular abnormalities in facioscapulohumeral muscular dystrophy. A general association with genetic and therapeutic implications, Brain, 110:631, 1987. 14. Kohl, S. et al., RDS/peripherin gene mutations are frequent causes of central retinal dystrophies, J. Med. Genet., 34:620, 1997. 15. Sunga, R.N. and Sloan, L.L., Pigmentary degeneration of the retina: early diagnosis and natural history, Invest. Ophthalmol., 6:309, 1967. 16. Sugiyama, F. et al., Concordance of murine quantitative trait loci for salt-induced hypertension with rat and human loci, Genomics, 71:70, 2001. 17. Kim, J.H. et al., Genetic analysis of a new mouse model for non-insulin dependent diabetes, Genomics, 74:273, 2001. 18. Ikeda, A. et al., Genetic modification of hearing in tubby mice: evidence for the existence of a major gene (moth1) which protects tubby mice from hearing loss, Hum. Mol. Genet., 8:1761, 1999. 19. Ikeda, A., Naggert, J.K., and Nishina, P.M., Genetic modification of retinal degeneration in tubby mice, Exp. Eye Res., submitted. 20. Williams, R.W. et al., Genetic and environmental control of variation in retinal ganglion cell number in mice, J. Neurosci., 16:7193, 1996. 21. Danciger, M. et al., A QTL on distal chromosome 3 that influences the severity of light-induced damage to mouse photoreceptors, Mamm. Genome, 11:422, 2000. 22. Cornall, R.J. et al., The generation of a library of PCR-analyzed microsatellite variants for genetic mapping of the mouse genome, Genomics, 10:874, 1991. 23. Silver, L.M., Mouse Genetics: Concepts and Applications, Oxford University Press, New York, 1995. 24. Darvasi, A. et al., Detecting marker-QTL linkage and estimating QTL gene effect and map location using a saturated genetic map, Genetics, 134:943, 1993. 25. Darvasi, A. and Soller, M., A simple method to calculate resolving power and confidence interval of QTL map location, Behav. Genet., 27:125, 1997. 26. Justice, M.J., Jenkins, N.A., and Copeland, N.G., Recombinant inbred mouse strains: models for disease study, Trends Biotechnol., 10:120, 1992. 27. Williams, R.W., Strom, R.C., and Goldowitz, D., Natural variation in neuron number in mice is linked to a major quantitative trait locus on Chr 11, J. Neurosci., 118:138, 1998. 28. Plomin, R. et al., Use of recombinant inbred strains to detect quantitative trait loci associated with behavior, Behav. Genet., 21:99, 1991. 29. Groot, P.C., et al., The recombinant congenic strains for analysis of multigenic traits: genetic composition, FASEB J., 6:2826, 1992. 30. Nadeau, J.H. et al., Analyzing complex genetic traits with chromosome substitution strains, Nat. Genet., 24:221, 2000. 31. Moore, K.J. and Nagle, D.L., Complex trait analysis in the mouse: the strengths, the limitations and the promise yet to come, Annu. Rev. Genet., 34:653, 2000.

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32. Manly, K.F. and Olson, J.M., Overview of QTL mapping software and introduction to map manager QT, Mamm. Genome, 10:327, 1999. 33. Churchill, G.A. and Doerge, R.W., Empirical threshold values for quantitative trait mapping, Genetics, 138:963, 1994. 34. Lander, E.S. et al., MAPMAKER: an interactive computer package for constructing primary genetic linkage maps of experimental and natural populations, Genomics, 1:174, 1987. 35. Lincoln, S.E., Daly, M.J., and Lander, E.S., Mapping genes controlling quantitative traits with MAPMAKER/QTL, Whitehead Institute Technical Reports, 2nd ed., 1992, available at http://wwwgenome.wi.mit.edu/ftp/distribution/software. 36. Basten, C., Weir, B.S., and Zeng, Z.-B., Zmap — A QTL cartographer, in Proceedings of the 5th World Congress on Genetics Applied to Livestock Production: Computing Strategies and Software, Smith, C. et al., Eds., 65, 1994. 37. Sen, S. and Churchill, G., A statistical framework for quantitative trait mapping, Genetics, 159:371, 2001. 38. Grupe, A. et al., In silico mapping of complex disease-related traits in mice, Science, 292:1915, 2001. 39. Paigen, K. and Eppig, J.T., A mouse phenome project, Mamm. Genome, 11:715, 2000. 40. Darvasi, A., Experimental strategies for the genetic dissection of complex traits in animal models, Nat. Genet., 18:19, 1998. 41. Flaherty, L., Congenic strains, in The Mouse in Biomedical Research, Foster, H.L., Small, J.D., and Fox, F.G., Eds., Academic Press, New York, 1981, 215. 42. Markel, P. et al., Theoretical and empirical issues for marker-assisted breeding of congenic mouse strains, Nat. Genet., 17:280, 1997. 43. Antoch, M.P. et al., Functional identification of the mouse circadian Clock gene by transgenic BAC rescue, Cell, 89:655, 1997. 44. Hamilton, B.A. et al., The vibrator mutation causes neurodegeneration via reduced expression of PITP alpha: positional complementation cloning and extragenic suppression, Neuron, 18:711, 1997. 45. Wilson, S.M. et al., A mutation in Rab27a causes the vesicle transport defects observed in ashen mice, Proc. Natl. Acad. Sci. U.S.A., 97:7933, 2000. 46. Naggert, J.K. et al., Hyperproinsulinaemia in obese fat/fat mice associated with a carboxypeptidase E mutation which reduces enzyme activity, Nat. Genet., 10:135, 1995. 47. Patil, N. et al., A potassium channel mutation in weaver mice implicates membrane excitability in granule cell differentiation, Nat. Genet., 11:126, 1995. 48. Daiger, S.P. et al., Data services and software for identifying genes and mutations causing retinal degeneration, Invest. Ophthalmol. Vis. Sci., 39:S295, 1998. 49. Lovett, M., Kere, J., and Hinton, L.M., Direct selection: a method for the isolation of cDNAs encoded by large genomic regions, Proc. Natl. Acad. Sci. U.S.A., 88:9628, 1991. 50. Duyk, G.M. et al., Exon trapping: a genetic screen to identify candidate transcribed sequences in cloned mammalian genomic DNA, Proc. Natl. Acad. Sci. U.S.A., 188:8995, 1990. 51. Schuler, G.D. et al., A gene map of the human genome, Science, 274:540, 1996. 52. Altschul, S.F. et al., Basic local alignment search tool, J. Mol. Biol., 215:403, 1990. 53. Burset, M. and Guigo, R., Evaluation of gene structure prediction programs, Genomics, 34:353, 1996. 54. Ausubel, F.M. et al., Current Protocols in Molecular Biology, Wiley, New York, 2001.

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Mutagenesis and Genetic Screens in the Mouse Timothy P. O’Brien

CONTENTS Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .93 Building the Mouse Mutant Resource . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .93 Mutagenesis in the Mouse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .95 Whole Animal ENU Mutagenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .95 Chemical Mutagenesis of Mouse Embyronic Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . .96 Genetic Screening Strategies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .97 Genome-Wide Dominant and Recessive Screens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .97 Screening for Recessive Mutations at a Single Locus . . . . . . . . . . . . . . . . . . . . . . . . . . . . .100 Screening for Recessive Mutations in a Defined Genomic Region . . . . . . . . . . . . . . . . . . .100 Screening for Modifiers and Interactive Loci . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .104 Sequence-Based Analysis and Chemical Mutagenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . .105 The Future of Mutagenesis and Genetic Screens in the Mouse . . . . . . . . . . . . . . . . . . . . . . . . . .105 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .106

INTRODUCTION Heritable mutations are critical for functional genetic analysis. A variety of approaches are available for building the collection of genetic variants in the mouse. These include the detection of spontaneous mutations, introducing alterations using transgenics, gene targeting, insertional mutagenesis or gene trapping, and treatment with chemicals or radiation to induce genetic lesions. This chapter focuses on methods for recovering genetic variants in the mouse using chemical mutagenesis and phenotype-driven screens. Phenotype-driven screens can be performed without prior knowledge of the gene and represent an unbiased survey of genomic content and molecular activity. In the mouse the chemical N-ethyl-N-nitrosourea (ENU) induces point mutations at a high rate, permitting efficient systematic phenotype-based surveys of gene function. Basic principles of mouse and embryonic stem (ES) cell mutagenesis and various breeding strategies are introduced to serve as a guide for performing screens and to provide an appreciation of mouse models generated using this approach.

BUILDING THE MOUSE MUTANT RESOURCE Several model organisms from bacteria and yeast to Caenorabditis elegans, Drosophila, and zebrafish have demonstrated the utility of genetic analysis to reveal the nature and complexity of gene function. In mammals, the mouse delivers this power of genetics. The mouse offers many well-characterized inbred strains that are genetically diverse, the ability to specifically manipulate or randomly mutagenize the genome, and comprehensive characterization of developmental and physiological pathways for defining functional similarity with humans. High-resolution genetic maps, physical maps, and genomic sequence shared between mouse and human allow for comparison and exchange 0-8493-0864-X/02/$0.00+$1.50 © 2002 by CRC Press LLC

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of information about gene function. In combination, these tools and resources underscore the capacity of the mouse to provide models for understanding human development, disease, and disability. Important insights have been gained from the analysis of a relatively small number of mouse models.1,2 For the eye, the Mouse Locus Catalog (MLC) lists ~130 loci associated with abnormal development, structure and function (http://www.informatics.jax.org/searches/noforms_mlc_omim.cgi). Although significant, this collection does not account for the number of genes and genetic interactions required to construct and maintain the performance of an organ as complex as the eye or to model the number of genes that play a significant role in human eye disorders. Thus, progress depends on efforts to fill gaps in existing data sets and resources. Whole-genome sequencing and functional genomic approaches, such as gene expression arrays and mutagenesis screens, hold the promise for narrowing this gap to achieve completeness and integration in biology.3-6 Analysis of mammalian genetic content and organization has been transformed by the availability of whole-genome sequences for human, mouse (in progress), and several other model organisms.7-11 One of the emerging challenges is to link an estimated 35,000 to 45,000 genes in mammals with specific biological functions. Classification of genes according to functional motifs using sequence comparisons, examination of expression patterns and profiles, biochemical studies, and mutational analyses are valuable approaches for assigning gene function. The insights afforded by the current collection of mouse mutants and the impact of the large sets of genetic variants in other model organisms has motivated the goal of expanding the mouse mutant resource to include variant alleles for all of the genes.12 Mutations have been recovered or engineered for a modest percentage of the genes in the mouse, and many of these genes are represented by just a single mutant allele. Banks of ES cells harboring engineered disruptions in all of the genes are under construction (http://tikus.gsf.de/), (http://socrates.berkeley.edu/~skarnes/). However, a set of complete loss-of-function or knockout mutations is only the beginning. There is a growing appreciation for the number of alternative transcripts expressed by a single locus, the degree of post-translational modification of the encoded protein, and the diverse roles played by a given gene over the life span of an organism. To address this diversity, gene targeting technologies have evolved to generate conditional, tissue specific, and partial loss-of-function alleles in an effort to examine the full range of gene activity.13,14 Chemical mutagenesis using ENU combined with phenotype-based screens offers a powerful and complementary approach for annotating gene function.15-20 Extensive mutational access to a locus will be crucial for identifying mouse models that represent inactivating as well as subtle mutations that underlie human disease. As a point mutagen ENU can induce a series of mutations leading to amorphic/null (loss-of-function), hypomorphic (partial loss-of-function), antimorphic (opposing or dominant negative function), and hypermorphic (gain-of-function) phenotypes. The function of a number of genes has been revealed using ENU mutagenesis in the mouse.21,22 In notable instances, as shown at the embryonic ectoderm development (eed) locus or the quaking (qk) locus, a series of alleles was critical for uncovering a range of gene functions.23,24 Another significant feature is that ENU often induces missense mutations. Deleterious changes in amino acid sequence lead to the identification of key residues required to configure functional motifs or whose modification is critical for activity. Therefore, ENU-induced mutations will also serve as valuable reagents for studies relating protein structure and function.25 Networks of genetic interactions guide biological processes. Gene products act as part of multimeric protein complexes and participate in intricate biochemical, physiological, and developmental pathways. One attractive application of ENU mutagenesis is to extend the mouse mutant resource while dissecting networks of gene interactions.18 This is accomplished through sensitized screens that take advantage of the existing collection of induced or engineered mutants. Genetic screens can be designed to uncover mutations that modify the original phenotype, revealing enhancer or suppressor loci. These screens can also identify mutations that present phenotypes resulting from non-allelic non-complementation, where the trait is exhibited in a mouse that is heterozygous for a mutated

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allele at two independent loci. The high degree of genetic complexity suggests that such screens could be very productive for gaining insights into the molecular mechanisms controlling specific biological processes.

MUTAGENESIS IN THE MOUSE The goal of mutagenesis for functional genetic studies is to increase the frequency and variety of mutations that will be recovered in a phenotype-driven screen. Over hundreds of years mouse fanciers and geneticists have collected spontaneous mutations essentially using phenotype-based screens. These classical mutants present phenotypes that are readily detected as changes in coat color, morphology, or behavior and have contributed greatly to genetic studies of developmental and neurological disorders.26 However, spontaneous mutations are too rare to be useful for building the mouse mutant resource using systematic phenotype screens. Studies that have involved the phenotypic assessment of large numbers of mice and historical control groups from mouse genetics research institutes, such as The Jackson Laboratory (TJL), Oak Ridge National Laboratory (ORNL), and the Medical Research Council (MRC), Mammalian Genetics Unit, Harwell, have been used to document the incidence of spontaneous mutations in mice.27-29 Generally, mutations measured at visible specific coat-color loci occur at a rate of 1 mutation per locus in every 100,000 offspring (1 × 10-5/locus/gamete). The effects of radiation and various chemical agents on the mammalian germ line have been assayed in experiments conducted at the MRC Radiobiology Unit, Harwell and ORNL. These studies used the specific locus test (SLT), a screening protocol that detects mutations induced in seven recessive visible marker loci.30 These seven loci, six altering coat color and one ear morphology, are carried in T-stock (tester) mice. In the classic SLT, male mice are exposed to mutagens and crossed to T-stock females. The offspring of this cross typically do not exhibit a phenotype since the original T-stock mutations are recessive. However, if a new mutation has been induced at any of the seven specific tester loci, the offspring will present the recessive coat color or short-eared phenotype of the corresponding original mutation. The SLT has been used to characterize a variety of mutagens, such as radiation or the chemical agent chlorambucil, that often induce large chromosomal deletions and rearrangements in mice.31,32 The SLT also established that ENU is an extremely powerful germ line mutagen capable of inducing a mutation at a specific locus in 1 of every 667 gametes screened (1.5 × 10-3/locus/gamete).33,34

WHOLE ANIMAL ENU MUTAGENESIS ENU acts as an alkylating agent by directly transferring an ethyl group to oxygen or nitrogen radicals in DNA. Point mutations are the result of base-pair substitutions owing to mispairing of the ethylated base during DNA replication.35 In mice, modifications most frequently occur at A/T base pairs, with 44% A/T → T/A transversions and 38% A/T → G/C transitions documented through sequence analysis of 62 ENU-induced alleles from 24 genes.36 Changes involving additional base-pairs and small deletions have also been observed. The base-pair changes cause differing effects at the locus, with a reported 26% altering splicing, 10% resulting in nonsense mutations, and 26% resulting in missense mutations of the translated protein.36 This spectrum of molecular lesions underlies the mutational capacity of ENU to generate a variety of dysfunctional alleles at a single locus. In the mouse, ENU most effectively mutagenizes spermatogonial stem cell populations. Effective mutagenic doses result in a ~10- to 20-week sterility period because of the depletion of spermatogonial stem cells in the testes. Surviving spermatogonial stem cells give rise to clonal populations of mutagenized sperm that can be sampled in the offspring (commonly referred to as the G1 progeny) of the mutagenized male (designated as G0). Because the populations of sperm are clonal,

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the number of mutagenized sperm (gametes) sampled should be monitored to avoid the isolation of repeat or “cluster” mutations. Generally, collecting 30 to 50 G1 offspring per ENU-treated G0 male is considered to provide adequate sampling while maintaining the likelihood that uniquely mutagenized genomes are being screened.37 Optimal ENU doses in mice reflect a balance between the mutagenic and toxic effects of the treatment.38,39 Administration of overall higher doses of ENU increases the mutation frequency. However, doses that are too high can cause permanent sterility, death, or lead to death almost immediately following fertility return. Effective ENU doses are determined on the basis of the percentage of mice that return to fertility, the length of the sterility period, and the ability to produce adequate numbers of G1 offspring. Several doses of ENU have been shown to be mutagenic in mice, with standard treatments in the range of 200 to 400 mg/kg of body weight administered in fractionated doses of 100 mg/kg per weekly injection. The highest doses, such as 4 × 100 mg/kg ENU, are tolerated by some F1 hybrid mice, while other hybrid and most inbred strains can only tolerate doses in the 2 × 100 to 3 × 100 mg/kg range. Although the optimal doses for several inbred and hybrid strains of mice have been reported,39,40 it is recommended that a range of doses be used when initiating an ENU mutagenesis experiment in mice. Ultimately, the effectiveness of ENU treatments is revealed by the recovery of heritable mutations at a high frequency. Mutation rate is dependent on a number of variables. Dosage, treatment regimen, selection of inbred mouse strain or hybrid combination, dominant vs. recessive screen, and assays used to detect phenotypes are all factors in projecting a mutation rate. In addition, gene size and functional composition influence mutation recovery at a specific locus, with the trend that larger genes are mutated with greater frequency. In mutagenesis screens it is generally considered that recessive mutations present phenotypes much more often than do dominant mutations. A comparison of ENU-induced dominant cataract and recessive specific-locus mutations indicated that the perlocus mutation rate to recessive alleles was six times greater than the per locus rate for dominant alleles.41 Several studies, including the SLT, have demonstrated the variability of ENU mutation rates at an individual locus, ranging from as high as 1 in 175 to lower than 1 in over 4000 gametes screened, while still other loci may be even more refractory to ENU mutagenesis.37,42 Overall, a mutation rate of 1 of every 1000 gametes screened (1 × 10-3/locus/gamete) for a gene of average mutability can be considered a guide for planning an ENU mutagenesis experiment.

CHEMICAL MUTAGENESIS OF MOUSE EMBRYONIC STEM CELLS Mouse ES cells can contribute to all tissues during development, including the germ line, permitting genetic modifications to be introduced in culture and analyzed in the mouse. Using mouse ES cells to combine advantages inherent to cell culture systems with chemical mutagenesis offers extraordinary possibilities for using phenotype-based and genotype-based approaches to address gene function.43 The ability to quality control, modulate, and readily assay mutation rate, higher mutation frequency, a wider spectrum of mutagens, shortened breeding strategies in mice, direct screening using ES cell culture, and archives of frozen mutagenized ES cell clones are attractive features of ES cell mutagenesis. Effective mutagenesis of ES cells using the point mutagens ENU and ethylmethanesulfonate (EMS, another alkylating agent commonly used in flies and worms), as well as the frameshift mutagen ICR191, has been demonstrated.43-45 Mutagenized ES cells retain germ line competence and screening pedigrees of mice derived from EMS-treated ES cells for visible abnormalities resulted in the identification of a range of heritable mutations. In addition, intercrosses of the G1 siblings from mutagenized ES cell germ line chimeras produced few if any offspring, suggesting that these mice carried a high load of recessive lethal mutations.44

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Mutation rates in ES cells are assayed by screening for loss-of-function mutations at the X-linked hypoxanthine phosphoribosyl transferase (Hprt) locus. Loss of Hprt function confers 6-thioguanine resistance allowing mutated clones of ES cells to be identified using drug selection. ENU and EMS induced mutations in ES cells at a frequency of approximately 1 in 1000 at the Hprt locus, a rate comparable to whole-animal ENU-induced recessive mutations. In addition, inhibition of O6-alkylguanine-alkyltransferase-mediated DNA repair using O6-benzyl-guanine increased the mutation rate at Hprt two- to fourfold.45 Sequence analysis of the Hprt alleles showed that base-pair substitutions resulting in missense and nonsense mutations predominated. The majority of ENU-induced mutations affected A/T base pairs whereas EMS primarily induced G/C → A/T transitions. The high mutation load, mutagen choice and fewer mice are significant advantages of the ES cell mutagenesis approach, while the production of chimeras is a significant consideration. However, the option to induce mutations in mice or in ES cells allows for alternative and complementary strategies and greatly enhances the opportunity to gain mutational access to build a more complete collection of mouse mutants.

GENETIC SCREENING STRATEGIES Inducing mutations is the first step in generating a mouse model for functional analysis. To isolate mutations, genetic screens are performed to provide mice that are suitable for phenotyping. A variety of breeding strategies and chromosomal reagents allow for screens that survey the entire genome as well as focus on specific genes, chromosomal regions, and areas of biology. In this section examples of basic strategies using the available genetic resources in mice are provided. As with other model organisms, growing collections of mutants and sets of chromosomal reagents combined with innovative phenotyping will lead to increasingly customized genetic screens in mice.

GENOME-WIDE DOMINANT AND RECESSIVE SCREENS A genome-wide screen for dominant mutations is simple and efficient. In this screen mutagenized G0 male mice are mated with wild-type females and the first generation (G1) offspring are screened for the desired phenotype (Figure 7.1). A collection of more than 200 dominant heritable mutations has been assembled in an extensive mutagenesis screen to detect defects in eye morphology. A total of 92 ENU-induced mutations, 56 of which have been mapped to 22 different loci, identified a variety of alleles for a number of genes that underlie phenotypes such as microphthalmia and cataracts in mice.46 A three-generation breeding strategy is used in a genome-wide screen for recessive mutations in mice (see Figure 7.1).15 In this screen, ENU-treated G0 males are mated with wild-type females to generate G1 offspring that are heterozygous for a fixed complement of induced mutations. The G1 mice are used to establish families (pedigrees) of mice. G1 males are mated with wild-type partners to produce G2 females, half of which are now heterozygous for a mutation at a given locus. However, it should be noted that all of the G2 offspring carry a subset of mutations derived from their G1 parent. The G2 females are backcrossed with their G1 fathers to produce G3 offspring that are homozygous for a fraction of the genome-wide mutations captured in the G1 founder. For a given locus only 1 /8 of the G3 progeny will carry a homozygous mutation. Screening of ~20 G3 offspring per G1 male provides strong confidence that all the mutations in the G1 founder have been surveyed for a recessive phenotype. Using mutagenized ES cells a genome-wide recessive screen can be performed in two generations (Figure 7.2).43 In this case the mutagenized genome of a diploid ES cell clone is considered to be fixed in the chimeric male. The mutagenized ES cell chimeric male is used to establish a pedigree and is mated to a wild-type female to produce G1 offspring. Random intercrossing of G1 siblings can

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ENU G0

X

X

*

G1

X

*

* G2 offspring:

or

* G3 offspring:

G1

* * *

*

dominant mutation

recessive mutation

or

*

FIGURE 7.1 Genetic screen for isolating genome-wide dominant and recessive mutations following whole-animal ENU mutagenesis. Mutations (*) induced in the G0 male are captured in the G1 offspring and these can be screened for a dominant phenotype. Three generations are required to isolate a recessive mutation. A backcross screening strategy following a mutation at a single locus is shown. G1 males are mated to produce G2 daughters that are either carriers of a mutation at a specific locus (top G2 female) or wildtype (bottom G2 female). G2 daughters are backcrossed to their G1 fathers. Only half of the matings (top G2 female) will result in G3 offspring that are homozygous for a mutation at a specific locus (G3 offspring shown on the right). In the other mating (bottom G2 female) none of the G3 offspring are homozygous for the mutation at the specific locus (shown on the left). Therefore, only one of eight G3 offspring reveals a recessive mutation at a given locus.

be used to render mutations homozygous. However, since matched pairs of heterozygous mutation carriers may not be selected, this cross is not assured to render a mutation homozygous at a specific locus. Alternatively, G1 females can be backcrossed to their chimeric father and in each instance one fourth of the progeny will carry a homozygous mutation at a given locus. A genome-wide recessive screen provides the opportunity to survey all the mutations induced throughout the genome. Dominant mutations as well as recessives will be present in the screening class progeny. In fact, it is important to consider that the progeny screened carry multiple mutations. The larger the number of phenotype screens performed, the more likely any one or more of these mutations will be detected. Several centers have been established to apply large-scale mutagenesis and

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ENU, EMS, ICR191, other ES cells

*

X

* G1 offspring:

recessive mutation

or

X

chimera

* *

* * * *

G2 offspring:

* * *

*

*

* or

chimera

recessive mutation

*

* *

*

FIGURE 7.2 Genetic screen to isolate genome-wide recessive mutations using mutagenized ES cells. Mutations (*) are captured in a chimeric male derived from a mutagenized ES cell clone. Two generations are required to isolate a recessive mutation. Chimeric males are mated to produce G1 offspring, each of these will carry a mutation at a specific locus that has been fixed in the chimeric male. In the scheme shown G1 daughters are backcrossed with their chimeric father to produce G2 offspring. All crosses result in G2 offspring that are homozygous for a mutation at a specific locus. Therefore, one of four G2 offspring reveals a recessive mutation at a given locus.

phenotyping to generate mouse models of human disease (Table 7.1). In these programs the mice are systematically screened using a battery of phenotypic assays.47-50 This has resulted in the recovery of mutants in a high number of G1 pedigrees.47 It is also possible that multiple mutations will lead to a phenotype that is complex, resulting from mutations in more than one gene. In a focused screen to recover recessive mutations that disrupt mouse embryogenesis at midgestation, 5 mutant lines were recovered in a screen of 86 lines, with 1 line that appeared to have a complex genetic basis.51 In general, extraneous mutations complicate genetic analysis; therefore, mutants derived from ENU mutagenesis screens should be backcrossed to a selected inbred strain to establish a genetically defined stock.

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TABLE 7.1 Mutagenesis and Phenotyping Centers for Mouse Models of Human Disease UK Mouse Genome Centre, Harwell - ENU Mutagenesis Programme..................(http://www.mgu.har.mrc.ac.uk/mutabase/) ENU-Mouse Mutagenesis Screen in the German Human Genome Project .......................(http://www.gsf.de/isg/groups/enu/) Neuroscience Mutagenesis Facility at The Jackson Laboratory ....(http://www.jax.org/resources/documents/nmf/index.html) Mouse Heart, Lung, Blood and Sleep Disorders Center at The Jackson Laboratory ......(http://www.jax.org/hlbs/index.html) Baylor College of Medicine Mouse Genome Project .............................(http://www.mouse-genome.bcm.tmc.edu/Home.asp) Tennessee Mouse Genome Consortium .....................................................................................................(http://tnmouse.org/) Mutagenesis at the McLaughlin Research Institute ...................... (http://www.montana.edu/wwwmri/mutants/protocol.html) Centre for Modeling Human Disease at the Samuel Lunenfeld Research Institute ..........................(http://cmhd.mshri.on.ca/)

SCREENING FOR RECESSIVE MUTATIONS AT A SINGLE LOCUS A series of alleles at a specific locus can be recovered in a single generation. The strategy for this screen is identical to that used in the SLT. A mutagenized male is crossed to a female that is homozygous for a recessive mutation at the test locus (Figure 7.3). The G1 offspring are screened for the desired phenotype; however, this may be more or less severe, or add or remove a feature of the original homozygous mutant phenotype. Thus, new alleles can be used to compare variations in function with the molecular nature of the series of mutations alone or in combination. Recessive mutations associated with embryonic or juvenile lethality and sterility can be isolated using linked markers and alternative breeding schemes (discussed below).

SCREENING FOR RECESSIVE MUTATIONS IN A DEFINED GENOMIC REGION An intensive investigation of the content and function of a defined genomic subinterval can be achieved in a regionally directed mutagenesis screen. Region-specific screens are facilitated by reagents such as large chromosomal deletions and inversions coupled with visible markers for tracking classes of progeny throughout the breeding scheme.15,16,52,53 Chromosomal aberration resources, such as sets of deficiencies covering ~70% of the genome, have been central to the success of Drosophila as a model organism for genetic analysis. Chromosomal aberration resources do exist in the mouse and, where available, these have been used to dissect the functional content of a defined genomic region. For example, in a classic two-generation breeding scheme using deletions at the albino locus over 4500 mutagenized gametes were screened to recover 31 mutations at 10 different loci, providing several alleles of genes required for survival and fitness in the mouse.37 Traditionally, chromosomal alterations have been generated and recovered in whole animals as was performed to build the radiation and chemically induced deletion complexes surrounding the visible marker loci used in the SLT. Innovations in methods for generating chromosome modifications using ES cells have made it possible to develop chromosomal deletion and inversion reagents throughout the mouse genome. In one approach a positive/negative selection cassette (neo-tk) is randomly introduced or targeted to a specific location in the genome. Following radiation, drug selection permits the recovery of ES cell clones that have deleted the neo-tk selection cassette (FIAU exposure kills the cells that have not deleted the neo-tk cassette and still express the selectable herpes simplex virus thymidine kinase (tk) marker). Sets of overlapping large and small deletions analogous to the deletion complexes of the SLT can be recovered and characterized in ES cells, and selected deletions can be used to generate mice.54,55 A bank of ES cells containing neo-tk selection cassettes positioned along each chromosome will allow deletion complexes to be developed throughout the genome (DELBank, http://lena.jax.org/~jcs/Delbank.html). Another approach involves chromosomal engineering using Cre-loxP technology.56 Large chromosomal deletions or inversions can be generated by targeting loxP sites to each end of the region to be modified. The rearrangements are

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FIGURE 7.3 Genetic screen to isolate multiple alleles at a single locus. An ENU-treated male (G0) is mated with a female that is homozygous for a viable mutation (a). New alleles (a*) that fail to complement the original mutation are recovered in the G1 offspring.

engineered and verified in ES cells and used to establish stocks of mice.57 A complementary set of libraries of pre-made gene-targeting insertion vectors has been constructed to facilitate genome-wide chromosome engineering.58 Combining the chromosomal deletions or inversions with visible markers enhances the utility of region-specific genetic screens. Marked classes of progeny result in more efficient detection, recovery, and maintenance of mutations. The most widely used markers have been mutations at endogenous loci that confer a viable and visible phenotype such as altered coat color. Transgene systems that take advantage of coat color genetics, such as the K14-Ag transgene, have been used to tag the rearrangements generated using Cre-loxP chromosome engineering.56 The K14-Ag transgene uses the keratin-14 promoter to drive the expression of Agouti and acts as a dominant marker by yellowing the coat of black mice.59 Another transgene that can serve as a dominant marker is the pCX-EGFP transgene. The pCX-EGFP transgene drives the ubiquitous expression of the enhanced green fluorescence protein (EGFP) permitting visualization of ES cells, pre- and postimplantation embryos, newborn pups, and adult mice.60,61 Thus, classes of progeny in a genetic screen can be distinguished at all stages of life. Dual labeling incorporating the fluorescent color variants (red, cyan, and yellow) provides multiple options and greater flexibility in the design of marked genetic screens. An EGFPbased two-color marker system and bank of marked ES cells is being developed to complement the genome-wide chromosomal aberration resources in mice (T. P. O’Brien, unpublished data). Screening using a deletion takes advantage of a functionally haploid portion of a chromosome. In a classic two-generation regionally directed screen the mutagenized male is mated with a wildtype female and the mutation-bearing G1 female is then mated with a male that is heterozygous for a large deletion (Figure 7.4). In the G2 offspring an ENU-induced mutation positioned opposite the deletion will reveal the recessive phenotype of a gene within the deleted region. The use of visible markers distinguishes all classes of G2 progeny. A recessive lethal mutation is identified by the absence of the specifically marked test class mouse among the G2 offspring and recovered in the marked carrier littermate. In a variation of this screen the mutagenized male is mated directly with a deletion heterozygous female. A visible marker can be used to identify the G1 offspring for phenotyping. Although mutations resulting in embryonic or juvenile lethality and sterility are not recovered, this screen requires only one generation. Large stretches of regional haploidy for some genomic regions, such as gene rich segments of chromosomes, may not be tolerated in mice (for example, heterozygosity for a large deletion is lethal). For these regions a chromosomal inversion can be used to conduct a marked three-generation

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FIGURE 7.4 Genetic screens to isolate recessive mutations using deletions and whole-animal mutagenesis. A. An ENU-treated male is mated with a female that is homozygous for a dominant visible marker (Q). G1 females carrying a mutation (*) and the marker (Q) are mated with heterozygous deletion (discontinuous line) males that carry a second dominant visible marker (Z). Four classes of marked G2 offspring are generated, two that are not valued in the screen that can be discarded (QZ and Q), the mutation carrier (Z), and the test class (*/deletion) that can be screened for recessive phenotypes or detected as a lethal missing class (unmarked). B. In this variation the ENU-treated male is mated directly with a female that is heterozygous for a deletion and carries a dominant visible marker (Z). Mutagenized gametes are sampled (*1, *2, *3, *4, …) in the G1 offspring. G1 offspring that do not carry the deletion (Z) can be discarded and the deletion bearing offspring (unmarked) can be screened for a viable recessive phenotype.

genetic screen (Figure 7.5). The inversion, like a balancer chromosome used in flies, serves to suppress recombination over a large interval, carries a dominant visible marker, and is lethal when homozygous. In this scheme the mutagenized male is mated with a female heterozygous for the marked inversion. The marked G1 male offspring are mated again with an inversion stock female carrying a second visible marker. The G2 offspring can be distinguished and selected for intercrosses or matings back with the G1 male. Recessive lethal mutations or mice for phenotyping along with carriers can be identified among the marked G3 progeny. By using ES cell mutagenesis, a region-specific recessive screen can be performed in one generation (Figure 7.6). In this scheme the mutagenized ES cell chimeric male is mated directly to a

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FIGURE 7.5 Genetic screen to isolate recessive mutations using a marked balancer chromosome. An ENUtreated male is mated with a female that is heterozygous for a recessive lethal inversion (solid black arrow) that is tagged with a dominant visible marker (Q). G1 males carrying a mutation (*) and the marked lethal inversion are mated with females that are heterozygous for the marked lethal inversion and carry a second dominant visible marker (Z). Three classes of marked G2 offspring are generated (inversion homozygotes are lethal), two classes are not valued in the screen and are discarded (QZ and Z), and the mutation carrier (Q). Mutation carriers are mated (G2 intercross or G2 female backcrossed with G1 father) to generate two classes of G3 offspring (inversion homozygotes are lethal), the mutation carrier opposite a large inversion that suppresses recombination (Q), and the test class (*/*) that can be screened for recessive phenotypes or detected as a lethal missing class (unmarked).

heterozygous deletion female. The use of visible markers distinguishes all four productive classes of G1 progeny, two recessive mutation-bearing test classes, and the corresponding mutation carriers. Because recessive mutations are confined to the genomic region, this screen takes full advantage of the high mutation load offered through ES cell mutagenesis. Region-specific screens and associated reagents complement and extend genome-wide screens for genetic analysis in mice. Region-specific screens allow for shortened breeding schemes and isolation of mutations that are already mapped to a defined position on a chromosome. The deletion panels can be used for fine-resolution genetic mapping, and chromosomal inversions afford efficient maintenance of mutations generated using either approach. Classes of mutagenized progeny can be

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FIGURE 7.6 Genetic screen to isolate recessive mutations using deletions and mutagenized ES cells. Mutations (*) are captured in a chimeric male derived from a mutagenized ES cell clone that is heterozygous for a dominant visible marker (Q). One generation is required to isolate recessive mutations. The chimeric male is mated with heterozygous deletion (discontinuous line) females that carry a second dominant visible marker (Z). Four classes of marked G1 offspring are generated, a test class (*/deletion) that is marked (Q) and its counterpart mutation carrier (QZ), and a test class (*/deletion) that is unmarked and its counterpart carrier (Z). Each test class can be screened for recessive phenotypes or detected as a lethal missing class.

identified using visible markers. Thus, recessive lethal mutations are recovered as a missing class and appropriate mice for screening can be selected saving costs associated with labor intensive or late onset phenotyping. Knowledge of the genetic content of a chromosomal interval available from complete genomic sequence can serve as a guide in the selection and analysis of the region to be functionally surveyed.

SCREENING FOR MODIFIERS AND INTERACTIVE LOCI In many instances crossing a mutation onto different inbred strains of mice changes the phenotype.62 Strain surveys to reveal naturally occurring modifiers have been used to gain insight into networks of gene interactions in mice. The isolation of modifiers showing semidominant effects suggests that it will be feasible to recover such enhancer or suppressor alleles using simple mutagenesis screens in the mouse.18 In other instances, genetic interactions have been revealed in mice that are heterozygous for mutations at two different loci, as a result of non-allelic non-complementation of the phenotype.63 A screen for interactive loci can be performed in a single generation (Figure 7.7). In the scheme shown, a male that is homozygous for a recessive mutation (which can be spontaneous, chemically induced, or engineered) is mutagenized. This mouse is mated with either a mutant or wild-type female to screen for dominant modifier and non-allelic non-complementing mutations induced at other loci throughout the genome. The original mutation serves as an entry point for the dissection of complex genetic interactions and biological pathways, providing a more complete view of collective gene function.

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FIGURE 7.7 Genetic screens to isolate modifiers or interactive loci. An ENU-treated male that is homozygous for a viable recessive mutation (a) is mated with a wildtype female to generate offspring that are heterozygous for (a) and screened for mutations at a second locus (b*) that fail to complement the original mutation (scheme shown on the left). The same mutagenized male can be mated with females that are homozygous for (a) to generate offspring that are also homozygous for (a) that can be screened for dominant mutations (M*) that modify the original phenotype (scheme shown on the right).

SEQUENCE-BASED ANALYSIS AND CHEMICAL MUTAGENESIS Another approach for using random mutagenesis to investigate gene function is to screen directly for sequence alterations in the gene of interest. Estimates of mutation frequency have projected that ENU-induced lesions from a fractionated-dose whole-animal treatment protocol are on the order of 1 sequence change per 105 bp.64 In ES cells direct sequence analysis at the Hprt locus showed ~1 mutation of every 200 mutagenized clones. A bank of mutagenized ES cells representing a series of alleles in all the genes could be screened to identify those clones containing the mutation of interest.43 A variety of techniques including gel-based single-strand conformation polymorphism (SSCP) analysis, physical separation using denaturing high-pressure liquid chromatography (DHPLC), direct sequence analysis, and high-density oligonucleotide microarrays are envisioned for detecting ENU-induced base-pair changes in DNA. Once the mutation is identified using a combined genebased random mutagenesis approach, the phenotypic consequences can be analyzed in mice.

THE FUTURE OF MUTAGENESIS AND GENETIC SCREENS IN THE MOUSE The value of systematic mutagenesis screens has been clearly demonstrated in model organisms.65-68 Large-scale projects in these models have illustrated that even with the extraordinary numbers of genomes screened and mutants collected the genetic diversity of these organisms has not been fully appreciated. In these organisms focused screens continue to provide biological insights. In the mouse, phenotype-driven mutagenesis screens provide the opportunity to link an estimated 35,000 to 45,000 genes with their biological functions. Several centers have been established to conduct systematic large-scale screens combining random mutagenesis and high-throughput phenotyping (see Table 7.1). These centers will generate hundreds of mouse models for the study of human disease and disability. In addition, several technologies and reagents are being developed that will enhance the application of

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genome-wide and directed mutagenesis screens. Access to improved resources and approaches will increase the ease of performing mutagenesis screens in smaller programs or individual laboratories with extensive experience and expertise in focused areas of biology. A combination of large-scale programs and focused studies that use imaginative mutagenesis, genetic screening, and phenotyping strategies will be necessary to appreciate the genetic complexity of mammalian gene function.

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26. Doolittle, D.P. et al., Catalog of Mutant Genes and Polymorphic Loci, 3rd ed., Vol. 1, Genetic Variants and Strains of the Laboratory Mouse, Lyon, M.F., Rastan, S., and Brown, S.D.M., Eds., Oxford University Press, New York, 1996. 27. Russell, L.B. and Russell, W.L., Frequency and nature of specific-locus mutations induced in female mice by radiations and chemicals: a review, Mutat. Res., 296:107, 1992. 28. Schlager, G. and Dickie, M.M., Natural mutation rates in the house mouse. Estimates for five specific loci and dominant mutations, Mutat. Res., 11:89, 1971. 29. Favor, J., International Commission for Protection against Environmental Mutagens and Carcinogens. Working paper no. 4. Spontaneous mutations in germ cells of the mouse: estimates of mutation frequencies and a molecular characterization of mutagenic events, Mutat. Res., 304:107, 1994. 30. Davis, A.P. and Justice, M.J., An Oak Ridge legacy: the specific locus test and its role in mouse mutagenesis, Genetics, 148:7, 1998. 31. Russell, W.L., X-ray induced mutations in mice, Cold Spring Harbor Symp. Quant. Biol., 16:327, 1951. 32. Russell, L.B. et al., Chlorambucil effectively induces deletion mutations in mouse germ cells, Proc. Natl. Acad. Sci. U.S.A., 86:3704, 1989. 33. Russell, W.L. et al., Specific-locus test shows ethylnitrosourea to be the most potent mutagen in the mouse, Proc. Natl. Acad. Sci. U.S.A., 76:5818, 1979. 34. Hitotsumachi, S., Carpenter, D.A., and Russell, W.L., Dose-repetition increases the mutagenic effectiveness of N-ethyl-N-nitrosourea in mouse spermatogonia, Proc. Natl. Acad. Sci. U.S.A., 82:6619, 1985. 35. Noveroske, J.K., Weber, J.S., and Justice, M.J., The mutagenic action of N-ethyl-N-nitrosourea in the mouse, Mamm. Genome, 11:478, 2000. 36. Justice, M.J. et al., Mouse ENU mutagenesis, Hum. Mol. Genet., 8:1955, 1999. 37. Rinchik, E.M. and Carpenter, D.A., N-ethyl-N-nitrosourea mutagenesis of a 6- to 11-cM subregion of the Fah-Hbb interval of mouse chromosome 7: completed testing of 4557 gametes and deletion mapping and complementation analysis of 31 mutations, Genetics, 152:373, 1999. 38. Justice, M.J. et al., Effects of ENU dosage on mouse strains, Mamm. Genome, 11:484, 2000. 39. Weber, J.S., Salinger, A., and Justice, M.J., Optimal N-ethyl-N-nitrosourea (ENU) doses for inbred mouse strains, Genesis, 26:230, 2000. 40. Justice, M.J. et al., Effects of ENU dosage on mouse strains, Mamm. Genome, 11:484, 2000. 41. Favor, J., The frequency of dominant cataract and recessive specific-locus mutations in mice derived from 80 or 160 mg ethylnitrosourea per kg body weight treated spermatogonia, Mutat. Res., 162:69, 1986. 42. Shedlovsky, A. et al., Mouse models of human phenylketonuria, Genetics, 134:1205, 1993. 43. Chen, Y., Schimenti, J., and Magnuson, T., Toward the yeastification of mouse genetics: chemical mutagenesis of embryonic stem cells, Mamm. Genome, 11:598, 2000. 44. Munroe, R.J. et al., Mouse mutants from chemically mutagenized embryonic stem cells, Nat. Genet., 24:318, 2000. 45. Chen, Y. et al., Genotype-based screen for ENU-induced mutations in mouse embryonic stem cells, Nat. Genet., 24:314, 2000. 46. Favor, J. and Neuhauser-Klaus, A., Saturation mutagenesis for dominant eye morphological defects in the mouse Mus musculus, Mamm. Genome, 11:520, 2000. 47. Hrabe de Angelis, M.H. et al., Genome-wide, large-scale production of mutant mice by ENU mutagenesis, Nat. Genet., 25:444, 2000. 48. Nolan, P.M. et al., A systematic, genome-wide, phenotype-driven mutagenesis programme for gene function studies in the mouse, Nat. Genet., 25:440, 2000. 49. Nolan, P.M. et al., Implementation of a large-scale ENU mutagenesis program: towards increasing the mouse mutant resource, Mamm. Genome,11:500, 2000. 50. Soewarto, D. et al., The large-scale Munich ENU-mouse-mutagenesis screen, Mamm. Genome, 11:507, 2000. 51. Kasarskis, A., Manova, K., and Anderson, K.V., A phenotype-based screen for embryonic lethal mutations in the mouse, Proc. Natl. Acad. Sci. U.S.A., 95:7485, 1998. 52. Justice, M.J. et al., Using targeted large deletions and high-efficiency N-ethyl-N-nitrosourea mutagenesis for functional analyses of the mammalian genome, Methods, 13:423, 1997.

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53. Rinchik, E.M., Developing genetic reagents to facilitate recovery, analysis, and maintenance of mouse mutations, Mamm. Genome, 11:489, 2000. 54. You, Y. et al., Chromosomal deletion complexes in mice by radiation of embryonic stem cells, Nat. Genet., 15:285, 1997. 55. Thomas, J.W., LaMantia, C., and Magnuson, T., X-ray-induced mutations in mouse embryonic stem cells, Proc. Natl. Acad. Sci. U.S.A., 95:1114, 1998. 56. Mills, A.A. and Bradley, A., From mouse to man: generating megabase chromosome rearrangements, Trends Genet., 17:331, 2001. 57. Zheng, B. et al., Engineering a mouse balancer chromosome, Nat. Genet., 22:375, 1999. 58. Zheng, B., Mills, A.A., and Bradley, A., A system for rapid generation of coat color-tagged knockouts and defined chromosomal rearrangements in mice, Nucleic Acids Res., 27:2354, 1999. 59. Kucera, G.T., Bortner, D.M., and Rosenberg, M.P., Overexpression of an Agouti cDNA in the skin of transgenic mice recapitulates dominant coat color phenotypes of spontaneous mutants, Dev. Biol., 173:162, 1996. 60. Okabe, M. et al., ‘Green mice’ as a source of ubiquitous green cells, FEBS Lett., 407:313–319, 1997. 61. Hadjantonakis, A.K., Generating green fluorescent mice by germline transmission of green fluorescent ES cells, Mech. Dev., 76:79, 1998. 62. Rozmahel, R. et al., Modulation of disease severity in cystic fibrosis transmembrane conductance regulator deficient mice by a secondary genetic factor, Nat. Genet., 12:280, 1996. 63. Rancourt, D.E., Tsuzuki, T., and Capecchi, M.R., Genetic interaction between hoxb-5 and hoxb-6 is revealed by nonallelic noncomplementation, Genes Dev., 9:108, 1995. 64. Beier, D.R., Sequence-based analysis of mutagenized mice, Mamm. Genome, 11:594, 2000. 65. Nusslein-Volhard, C. and Wieschaus, E., Mutations affecting segment number and polarity in Drosophila, Nature, 287:795, 1980. 66. Kuwabara, P.E., Worming your way through the genome, Trends Genet., 13:455, 1997. 67. Haffter, P. et al., The identification of genes with unique and essential functions in the development of the zebrafish, Danio rerio, Development, 123:1, 1996. 68. Driever, W. et al., A genetic screen for mutations affecting embryogenesis in zebrafish, Development, 123:37, 1996.

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Section III Regional Ocular Pathology Genetic engineering and chemical mutagenesis are producing a plethora of novel phenotypes in mice. With completion of the mouse and human genome projects, it will be possible to better understand molecular mechanisms and pathways that affect disease. Identification and description of modifier genes, a difficult task in humans, will assume increasing importance in the near future. The close homology between mouse and human genomes means that mice play a critical role in both understanding and developing novel treatments for human disease. The aim of this section (Chapters 8 through 11) is to describe the regional pathology of the eye, using specific examples. Our goal is to build on the previous descriptions of normal anatomy (Chapters 1 through 3) and to describe the range of morphological variation that occurs in a specific tissue or structure to provide a framework for interpretation of new mutations. The simple structure of the lens will serve as an example of this approach. The lens consists of lens capsule, epithelium, and cortex—the latter almost totally acellular. The capsule can thicken or rupture. The epithelium can die, proliferate, or undergo metaplasia. The cortex can dissolve or abnormal cortex can be produced by altered lens epithelium. Many variations on these simple morphological themes occur as a result of alterations in different genes, but the underlying histopathological patterns remain constant. The value of presenting all of the reported cataract phenotypes is limited. However, knowledge of morphological mechanisms provides a sound basis for understanding novel histopathological findings. Because some diseases (e.g., glaucoma) involve multiple ocular structures, they will be discussed in detail in a single section, rather than being fragmented in several chapters. However, some material is discussed in more than one location, in order to illustrate a specific topic.

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The Anterior Segment Richard S. Smith, John P. Sundberg, and Simon W. M. John

CONTENTS Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .111 Lids and Ocular Adnexae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .112 Lids and Cilia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .112 Meibomian Glands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .113 Tarsal Glands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .115 Cornea and Sclera . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .116 Developmental Abnormalities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .116 Central Corneal Opacities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .116 Generalized Cornea Haze . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .117 Corneal Hyperplasia and Choristomas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .118 Stromal Hypoplasia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .118 Ocular Surface Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .120 Ocular Trauma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .122 Open Eyelids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .122 Infectious Disease—General Morphology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .124 Dystrophic Calcification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .124 Corneal Neovascularization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .125 Keratoplasty . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .127 Corneal Storage Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .127 Corneal Edema . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .130 Corneal Endothelial Proliferation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .132 Aqueous Outflow and Intraocular Pressure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .133 Classification of Glaucoma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .136 Developmental Glaucoma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .136 Open Angle Glaucoma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .136 Angle Closure Glaucoma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .136 Anterior Segment Findings in Glaucoma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .136 Posterior Segment Findings in Glaucoma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .139 Retina and Optic Nerve . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .139 Glaucoma and Glaucoma-Like Effects in Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .139 Age-Related Glaucoma in DBA/2J and Related Strains . . . . . . . . . . . . . . . . . . . . . . . . . . .141 Forkhead Transcription Factors and Glaucoma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .144 Microphthalmia and Anophthalmia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .148 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .151

INTRODUCTION This chapter reviews common histopathological lesions that occur in the lids and ocular adnexae, the cornea and sclera, and the aqueous production and drainage structures. The last includes the trabecular 0-8493-0864-X/02/$0.00+$1.50 © 2002 by CRC Press LLC

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meshwork, Schlemm’s canal, and the anterior uvea (iris and ciliary body), because of their intimate involvement in aqueous humor production and outflow. Since glaucoma and microphthalmia frequently involve anterior segment structures, the general histopathological features of both conditions are discussed in this chapter.

LIDS AND OCULAR ADNEXAE LIDS AND CILIA The external surface of the eyelids is covered by skin extending from the scalp and muzzle. Although there are many mutations that involve the skin and its associated structures,1,2 this discussion is limited to those that directly involve the eye and eyelids. Cilia are long, wide hair fibers produced by specialized follicles near the mucocutaneous junction, as described in Chapter 1. Microscopically, the pilosebaceous unit is identical to that of truncal hairs, but larger. Normal cilia curve outward, forming a tactile mesh in front of the globe that helps protect the cornea. If these hairs are malpositioned and directed toward the cornea (distichiasis; entropion) the corneal surface may be directly injured. The mucocutaneous junction of the eyelids, like other epithelial transition sites in the body, reflects a change of function. These are biologically active areas and often targets of specific pathogens. Ulceration can develop at this site in specific strains. Spontaneous, idiopathic ulcerative blepharitis develops in BALB/cJ, BALB/cByJ, CBA/J, and 129P3/J inbred strains of mice (Figures 8.1 and 8.4). The underlying abnormalities have not been defined. A Corynebacterium species has been isolated but the disease could not be reproduced in immunodeficient mice, which suggests that

FIGURE 8.1 Lid lesions. A. In balding (Dsg3bal) mice, a deficiency in desmoglein 3 is associated with ulcerative blepharitis (arrow). Original magnification × 50. B. At higher magnification, there is ulceration of the lid margin (arrow). Original magnification × 200. C. In the mouse mutation, tabby (EdaTa), there is complete absence of Meibomian glands. Original magnification × 100. D. A normal mouse has well-developed Meibomian glands (arrow). Original magnification × 100.

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this organism was secondary to an underlying abnormality 3-5 Ulcerative blepharitis is also a frequent finding in balding (Dsg3bal) and balding Pasteur (Dsg3bal-Pas) mutant mice, which have mutations of the desmoglein 3 gene (responsible for keratinocyte adhesion). Lack of desmoglein 3 is associated with formation of suprabasilar vesicles that are morphologically identical to those found in human and domestic animals with pemphigus vulgaris. There is usually a humoral-based autoimmune disease directed at desmoglein 3 in pemphigus vulgaris.6,7 While basal and squamous cell carcinoma of the eyelids is frequently identified in human eyes, neither variety has been reported in mice, although this may be due to a failure to examine the lids in transgenic mice that develop such lesions elsewhere. Ectropion causing secondary corneal damage occurs in human patients with lamellar ichthyosis.8 Mice develop several varieties of ichthyosiform disease, such as ichthyosis vulgaris,9 harlequin ichthyosis,10 and Netherton’s syndrome.11,12 Although not studied with regard to the lids and eyes, these mutant mice may represent potential models for human ocular disease.

MEIBOMIAN GLANDS The Meibomian glands are modified sebaceous glands that produce a lipid secretion as a component of the corneal tear film (see below).13,14 These large glands, like other holocrine glands, have a layer of flattened reserve cells at their periphery. The reserve cells differentiate into round cells with clear cytoplasm that contain small, round, clear vacuoles of uniform size. When the cells have matured, the secretion is extruded by cell rupture into a stratified squamous epithelial-lined duct that opens at the mucocutaneous junction. Abnormalities of these glands occur as (1) total loss of the structure, (2) hypoplasia of the gland, or (3) abnormalities of the duct (Table 8.1). The nonocular clinical findings are often more obvious, but eyelids are frequently involved. Meibomian glands are absent in three distinct spontaneous mouse Table 8.1 Mutant Mice with Eyelid Abnormalities

Mutation

Gene Symbol

Chr

Lesion

Ref.

Balding

Dsg3

18

Pemphigus-like, ulcerative blepharitis

6, 36

Crinkled

cr

13

Meibomian gland aplasia

1, 37, 38

Downless/Sleek

Edar

10


1, 39–41

Tabby

Eda

X


16, 23, 39

Asebia

Scd1

Meibomian gland hypoplasia, tarsal gland hypoplasia

1, 25

Hairless

hr

14

Meibomian ductal ectasia

1

Ornithine decarboxylase transgenic

Odc

NA

Meibomian ductal ectasia, hairless-like phenotype

33

Spermidine/Spermine N1–acetyltransferase transgenic

Sat

NA

Hairless-like phenotype

34

Bareskin

Bsk

11

Meibomian gland dystrophy and ductal ectasia

1, 2

Peach fuzz

Pfz

11

Meibomian gland dystrophy and ductal ectasia

35

NA = not applicable. Transgenes integrate into multiple sites throughout the genome. Integration sites vary with each line created even if the same construct is used.

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mutations, tabby (EdarTa), downless (Edardl), and crinkled (cr). that have similar ocular and systemic abnormalities (Figure 8.1; reviewed in Sundberg1,15 ). The clinical phenotype in these three mutations resembles the group of human diseases known as anhidrotic ectodermal dysplasia. Crinkled, downless, and tabby represent three distinct mutations, which have been mapped to different chromosomes and have been highly useful in understanding the ligand–receptor interactions.16-23 Hypoplasia or abnormal development of Meibomian glands occurs in three spontaneous allelic mutations that affect the stearoyl Coenzyme A desaturase1 (Scd 1) gene.1,24,25 The clinical phenotype was initially referred to as asebia because it was thought the mice lacked sebaceous glands.26 (Figures 8.1 and 8.2). However, sebaceous glands are present, but hypoplastic.27-29 All sebaceous glands associated with hair follicles, including those of the cilia and truncal hairs on the eyelids, are affected.25,28 Various degrees of mild sebaceous gland hypoplasia are also evident in mice lacking diacylglycerol acyltransferase (Dgat).30 Hypoplasia of the Meibomian glands is also seen in bareskin and peach fuzz mutant mice (see below). The third group of mutant mice with Meibomian gland abnormalities include those that have defects in the ducts leading from the glands. The hairless mouse mutation and the more severe allelic mutations at this locus including rhino mice are characterized by permanent loss of hair after completion of the first hair cycle.1,15 All these allelic mutant mice develop Meibomian gland ductal ectasia (Figure 8.2)31 Ornithine decarboxylase32,33 and spermidine/spermine N1–acetylspermidine transgenic mice develop cutaneous lesions similar to those found in rhino mice.34 Meibomian gland ectasia was observed in ornithine decarboxylase transgenic mice (Figure 8.3).33

FIGURE 8.2 Lid lesions. A. In a mouse homozygous for the asebia mutation (Scd1ab) there are no Meibomian glands in an area where they should be found (*). Original magnification × 100. B. In a normal mouse, Meibomian glands (M) are abundant and their cells filled with secretion (see also Figure 8.1D). Original magnification × 200. C. In asebia mice, a few typical Meibomian glands can be found (arrow), but they are not well developed. Original magnification × 200. D. Eyelids from a rhino mouse show highly folded epidermis (arrows) as well as large keratin-filled cysts (*). Original magnification × 20.

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FIGURE 8.3 Lid lesions. A. In a mouse with ornithine decarboxylase deficiency, there is massive wrinkling of the skin, similar to that seen in rhino mice. B. In an ornithine decarboxylase–deficient mouse, the epidermis of the lid is folded and there are large cystic structures (*) similar to those of rhino mice. Original magnification × 50. C. In a 32-day-old bareskin (Bsk/+) mouse there is hypoplasia of the Meibomian gland (arrow). Original magnification × 50. D. At higher magnification, there are only a few cells (arrow) that contain secretory material. Original magnification × 100.

TARSAL GLANDS Tarsal glands are typical sebaceous glands associated with the eyelid cilia. As discussed above, the three asebia mutations all have hypoplasia of the Meibomian and other pilosebaceous glands.15 Two strains of mice created by chemical mutagenesis have small but normal tarsal glands. The bareskin (Bsk) and peach fuzz (Pfz) mutant mice are due to autosomal dominant mutations.1,35 In both, the sebaceous duct becomes hyperplastic and fills with desquamated cornified cells (see Figure 8.3). Transmission electron microscopy reveals that sebocytes of the mutant mice contain vacuoles with electron opaque material that crystallizes when the cell ruptures. There is loss of cilia late in the disease. Both mutations map to mouse Chr 11 and may be allelic. Blepharitis may occur because of the excess sebum and keratin debris (Figure 8.4). The most common mutations in mice that involve the Meibomian and other adnexal glands are summarized in Table 8.1.

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FIGURE 8.4 Blepharitis and endophthalmitis. A. In an AKXD-23 mouse, a thin film of exudate covers the denuded periorbital epidermis (arrow). B. Dried exudate is present on the lids of a CBA/J mouse. C. Ulcerative blepharitis in a CBA/J mouse. To the left of the arrow there is hyperkeratosis at the mucoepidermal junction. Original magnification × 100. D. Higher magnification of C demonstrates loss of conjunctival epithelium (arrow) over a focus of acute inflammation. Original magnification × 200. E. Endophthalmitis in a C3H/HeJ mouse, an unusual complication of blepharitis. The cornea (C) and vitreous (V) are heavily infiltrated by neutrophils. The lens (L) has a diffuse cataract. Original magnification × 20. F. A higher power view of E illustrates the dense vitreous inflammatory infiltrate as well as acute retinal necrosis and detachment (arrow). Original magnification × 250.

CORNEA AND SCLERA DEVELOPMENTAL ABNORMALITIES Central Corneal Opacities The role of the Pax6 homeobox gene in development of the eye and especially of the lens is well known.42,43 Since the lens is a derivative of surface ectoderm, it is not surprising that the cornea is

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affected in Pax6 mutant mice. The gene is expressed in corneal and conjunctival epithelium both during development and in adults.44 Homozygosity for the gene mutation originally known as small eye (Sey, now Pax6Sey), a non-functional allele, results in severe ocular malformation with a small or absent lens and microphthalmia.45 Heterozygous Pax6Sey mice are less severely affected and demonstrate iris hypoplasia and central corneal opacities, comparable to those described in human aniridia or in Peters’ anomaly.46,47 Similar corneal opacities occur in several strains of inbred black mice as a low incidence background lesion.48,49 The corneal opacity is located centrally and the peripheral cornea is typically clear. A small clear spot is usually present near the center of the opacity (Figure 8.5). At early stages of lens formation (see Chapter 3) there is direct continuity between the corneal and lens epithelia. Normally, this connection disappears around E11, as the anterior chamber forms. The clinical appearance is due to persistence of this connection between corneal and lens epithelium, and the clear central spot is composed of epithelium. The aberrant intracorneal epithelium likely interrupts normal development of corneal stroma and Descemet’s membrane, producing irregular collagen deposition and corneal opacification. Occasionally, iris tissue may be trapped at the point of contact between lens and cornea, allowing iris vessels to invade the cornea, causing more extensive opacification. Although the lens is usually clear, there may be a dense cataract.

FIGURE 8.5 Developmental abnormalities. A. Persistence of lens stalk in E14 C57BL/6J embryo. The corneal and lens epithelia are continuous, interrupting the central corneal stroma that should be well developed by this age. Original magnification × 250. B. When the corneo-lenticular connection persists, it can be identified in adult mice by a central clear zone (arrow), surrounded by a corneal opacity of variable size.

Generalized Corneal Haze Rarely, inbred black mice develop diffuse corneal opacification accompanied by stromal neovascularization. A similar clinical phenotype in human eyes, termed autosomal dominant keratitis, was attributed to a mutation in the PAX6 exon 11 splice-acceptor site.50 Corneal opacification and vascularization are also found in aniridia, as a PAX6 human phenotype.51 Diffuse corneal stromal haze and neovascularization are also present at birth in mice homozygous for a targeted mutation of the LIM-homeodomain transcription factor LMX1B. Abnormalities are first detected at E15.5, as iris and ciliary body hypoplasia, a poorly formed anterior chamber, and loosely packed corneal stroma. At birth, the corneal stroma is thickened and there is vigorous stromal neovascularization. Electron microscopy demonstrates increased diameter of individual collagen fibers and an irregular arrangement of stromal collagen that account for the corneal haze (Figures 8.6 through 8.8).52

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FIGURE 8.6 Developmental abnormalities. Lmx1b+/+ and Lmx1b–/– newborn mice. The pupil (*) is irregular and the iris is hypoplastic compared to the normal mouse. (From Pressman, C., Genesis, 26:15–25, 2000. Reprinted by permission of Wiley-Liss, Inc., a subsidiary of John Wiley & Sons.)

Corneal Hyperplasia and Choristomas Corneal stromal and epithelial thickening with neovascularization characterizes the phenotype of mice with a construct that contains the mouse lens-specific αA-crystallin promoter linked to the cDNA encoding human fibroblast growth factor 7 (FGF7, formerly keratinocyte growth factor). The earliest morphological findings are present by E14.5 and there is progression through birth. These mice demonstrate enhanced cell division of the basal layers of the corneal epithelium that undergo hyperproliferation leading to abnormal differentiation with lacrimal gland formation in the corneal stroma.53 The cornea is not a normal location for the lacrimal gland and the aberrant tissue is known as a choristoma (in contrast to a hamartoma, which is a benign proliferation of tissue normal for a given location, but inappropriately arranged).31 It seems likely that this induced choristoma may stimulate neovascularization, because stromal implantation of certain cells is associated with vascular invasion.54,55 The epithelial hyperproliferation observed with FGF7 overexpression differs from that described in the corn1 mutation (see below).56 Stromal Hypoplasia Corneal stromal hypoplasia is less common, but has been reported in both transforming growth factor–β-2 (TGFβ-2) null mice57 and in transgenic mice with a 39-kb mouse proα1(II) collagen transgene with a deletion of exon 7 and intron 7 that alters the product of the Type II collagen gene (Col2a1).58 The corneal stroma of TGFβ-2 null mice at E18.5 is hypercellular and less than 25% of normal thickness (Figure 8.9). Because of multiple congenital defects, these mice die shortly after birth, so an adult phenotype is unknown. One explanation for the thin cornea would be a failure of collagen production, although ultrastructural studies have not been reported.57 In a different transgenic line, mutations in the Col2a1 gene were also associated with abnormal keratocytes and in addition demonstrated abnormal vitreous strands (Figure 8.9; see also Chapter 9).58

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FIGURE 8.7 Developmental abnormalities. A and B. Lmx1b+/+ and Lmx1b–/– newborn mice. The iris and ciliary body (boxed areas) demonstrate hypoplasia. Original magnification × 20. C and D. Enlargement of boxed areas indicating a shallow anterior chamber (*) and absence of ciliary folds (arrowhead). The iris stroma (arrows in C) is absent in the –/– mouse. Original magnification × 200. A to D Mallory’s trichrome stain. E and F. Location of corneal and lens epithelial defects (boxed areas) G and H. The corneal stroma (C) in the normal mouse is dense and avascular. The lens (L) epithelium is cuboidal. In the –/– mouse, the corneal stroma is disorganized and appears to contain less collagen. Several vascular channels containing erythrocytes (BC) are evident. The lens epithelium is columnar. Original magnification × 400. (From Pressman, C., Genesis, 26:15–25, 2000. Reprinted by permission of Wiley-Liss, Inc., a subsidiary of John Wiley & Sons.)

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FIGURE 8.8 Developmental abnormalities. Disturbed collagen fibrillogenesis in Lmx1b–/– mice. A. Collagen fibers (arrows) are arranged regularly in the +/+ mouse between keratocytes (K). B. Electron lucent spaces are found in the corneal stroma between keratocytes in the –/– mouse. A and B original magnification × 10,000. C. Higher magnification of boxed area in A demonstrates regular diameter of collagen fibers cut in cross section. D. In –/– mice, the collagen fiber diameter is greater than normal. C and D original magnification × 50,000. (From Pressman, C., Genesis, 26:15–25. 2000. Reprinted by permission of Wiley-Liss, Inc., a subsidiary of John Wiley & Sons.)

OCULAR SURFACE DISEASE Injury of the ocular surface by physical or genetic factors may have serious visual implications. The eyelids, corneal and conjunctival epithelium, and the complex tear film act as a functional physiological unit that plays a critical role in the health of the anterior segment. Maintenance of the tear film provides a smooth ocular surface for visual purposes and also acts as a barrier to invasion by microorganisms. Through blinking, the lids help to renew the tear film and spread it over the eye. In brief, the tear film consists of secretions of the Harderian, lacrimal, Meibomian, and accessory glands of the eyelid, as well as mucus provided by the conjunctival goblet cells. The corneal and conjunctival epithelial surfaces are also an important part of the surface barrier system.59,60 Damage to these structures or to the limbal stem cells leads to severe ocular consequences including stromal scarring, neovascularization, and corneal perforation.61,62 Lacrimal insufficiency is a common cause of ocular surface disease in human eyes.63 Mice homozygous for a nonfunctional Tgfβ1 gene develop focal lacrimal gland inflammation.64 In addition

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FIGURE 8.9 Eye defects in Tgfβ2–/– mice. A. Normal E18.5 eye. Inner (INL) and outer neuroblastic (ONL) layers and the optic nerve (ON) are indicated. B. Tgfβ2 null E18.5 eye. The inner neuroblastic layer is thickened and there is a thickened tunica vasculosa in the vitreous and attached to the lens (RVT). A and B original magnification × 50. Boxed area enlarged in C and D. C. +/+ mouse cornea with normal corneal epithelium (C) and stroma (S). D. The corneal epithelium is present in the –/– mouse, but the stroma is markedly thinner than normal. (A to D, from Doetschman, T., Development, 124:2659–2670, 1997. With permission.) E. C57BL/6 × DBA/2J mixed background normal control, E 18.5. The anterior chamber (*) is normal. F. Del3 transgenic line. The anterior chamber (*) is absent. G. Gly-85-1 transgenic line. The anterior chamber (*) is absent. H. In the normal mouse, the lamellar structure of the cornea is evident and keratocytes (arrow) are normal. I and J. In both Del3 and Gly-85-1 transgenic lines, the corneal stroma is disorganized with enlarged interfibrillar spaces and the keratocytes (arrows) are enlarged. E to G. Col2a1 mutant mice and controls. Original magnification × 50; H to J × 100. (From Ihanamaki, T., Eur. J. Ophthalmol., 6:427–435, 1996. With permission of Wichting Editore.)

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to systemic inflammation, dry eyes and blepharoconjunctivitis develop and are associated with lacrimal gland inflammation. The inhibition of both systemic and ocular effects by fibronectin peptides suggests a direct relationship between the disease and the absent protein.23 The corneal and conjunctival epithelia form a physical barrier to infection that few organisms can penetrate as long as it is unbroken. Once epithelial breakdown occurs, it is often associated with infection and corneal ulceration in humans and in older mice. Mice lacking the keratin 12 (Krt1-12) gene have fragile corneal epithelium that is easily damaged by minimal trauma (see below), although corneal infections were not observed in the 1-week period after epithelial removal. Based on the analogous clinical situation in human eyes, it is likely that a longitudinal study of Krt1-12–/– mice would demonstrate an increased incidence of keratitis. The number of desmosomes and hemidesmosomes was normal, eliminating this as an explanation for the epithelial fragility and suggesting that these mice are not a model for the recurrent corneal erosion syndrome in human eyes.65,66 The consequences of persistent epithelial loss are demonstrated by the morphological changes that occur in plasminogen-deficient (Plg–/–) mice.67 Control mice subjected to partial removal of corneal epithelium healed in seven days with clear corneas. The healing process in Plg–/– mice required up to three times as long and was accompanied by prolonged inflammation, corneal scarring, and neovascularization. Infection was avoided by administration of topical antibiotics, but would have been likely without them as a consequence of experimental disruption of the epithelial barrier.

OCULAR TRAUMA Even with excellent animal husbandry, mice inevitably come in contact with particles of bedding, potentially contaminated with urine and feces that can disrupt integrity of the tear film and corneal surface. In light of this fact it is surprising that infectious corneal disease is a rare event. Examination of large groups of mice makes it obvious that small bedding particles are often seen on the anterior corneal surface. These are usually uneventfully washed away by tears and blinking, but occasionally become embedded in the cornea. When that occurs, inflammatory cells quickly infiltrate the cornea. The severity of the inflammation depends on the extent of injury and whether it is accompanied by bacterial invasion. The foreign material is usually easy to identify with polarizing filters because of its anisotropy. Macrophages and giant cells often surround the foreign body (Figure 8.10). When sequestered in this way, the inflammation usually subsides leaving a residual corneal scar that can be quite extensive. In an inactive scar a foreign body can be demonstrated only with serial sections. Deeper injuries can rupture Descemet’s membrane. A combination of background disease8 and bedding-related ocular trauma likely explains many of the postnatally acquired opacities reported in C57BL/6 mice.68

OPEN EYELIDS As discussed earlier (Chapters 1, 3, and 4), open eyelids at birth result when the normal fusion of the eyelids around E15 fails to occur. The lids themselves are morphologically normal; the problem lies with exposure of the eyes as soon as the mouse is born. Direct corneal trauma from bedding and debris or from parents and littermates causes a break in the barrier to infection. This is complicated by persistence of induced epithelial defects because of drying, since normal blinking and spreading of tears is absent at birth and its lack accelerates tear film evaporation. Severe keratitis is frequent in mice with open lids at birth and is usually followed by dense corneal scarring and neovascularization similar to that seen in other corneal ulcers (see Figure 8.10). Perforation of the globe from liquefaction of the corneal stroma may occur.61

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FIGURE 8.10 Corneal inflammation. A. Corneal ulcer of unknown etiology in a CBA/J mouse. Loss of the corneal epithelium (arrow) provides a point of entry for bacteria. A dense inflammatory infiltrate extends from this point into more peripheral cornea. Original magnification × 100. B. At higher magnification, there is a dense infiltrate of neutrophils and lymphocytes. The posterior stroma is relatively unaffected. Original magnification × 250. C. In a chronic corneal ulcer, the corneal epitheium is thickened and the surface is becoming cornified (arrow). A mucoid exudate covers the base of the ulcer (*). Original magnification × 100. D. Both focal and diffuse keratitis (arrowhead) can produce a chemotactic response with a sterile accumulation of neutrophils in the anterior chamber—a hypopyon (arrow). This may be visible clinically, but does not necessarily indicate intraocular infection. Original magnification × 200. E and F. Occasionally, an inflammatory stimulus may elicit a fibrovascular proliferative response, termed a pyogenic granuloma. This is characterized by numerous large vascular channels (arrow) and by fibrous proliferation (*). Original magnification E × 200, F × 400. G. Central intracorneal foreign body in DBA/2J mouse. The foreign body, likely a piece of bedding, is associated with a mild chronic inflammatory infiltrate. Original magnification × 100. H. At higher magnification, lymphocytes, plasma cells, and new-formed blood vessels surround the foreign body. In addition foreign body giant cells are present (arrow). Original magnification × 400.

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INFECTIOUS DISEASE—GENERAL MORPHOLOGY Infectious corneal ulceration may be associated with ocular surface disease, trauma, exposure, and blepharoconjunctivitis. In the acute phase of infection, the stromal infiltrate consists largely of neutrophils. The agents that cause the corneal ulceration may exert a strong chemotactic effect that not only attracts leukocytes into the corneal stroma, but may also attract large numbers from iris vessels to the anterior chamber. When this occurs, leukocytes may accumulate in a concentrated layer in the inferior chamber, which is referred to as a hypopyon (see Figure 8.10). Bacterial colonies may be identified in the ulcerated epithelium or stroma and may invade the eye, causing endophthalmitis (see Figure 8.4). Depending on the organism, necrosis of the corneal stroma and Descemet’s membrane can lead to perforation and partial loss of intraocular contents as well as panophthalmitis (inflammation of all ocular layers) and secondary orbital cellulitis (diffuse orbital inflammation). Systemic sepsis may follow, although this is unusual. Less intense stimuli, such as low-grade blepharitis, tend to produce a chronic mild keratitis characterized by lymphocytes and plasma cells (see Figure 8.10). Stromal scarring and neovascularization are common sequelae. Herpes simplex keratitis can be induced in mice, and this system is used to model human herpes simplex infections, both primary and recurrent69 and to study specific defense mechanisms.70,71 In primary infections both linear epithelial ulceration (dendrites) and diffuse stromal opacification occur. The stromal haze increases in recurrent herpes and is often accompanied by corneal neovascularization. In recurrent disease, the stromal infiltrate is relatively sparse and composed of lymphocytes.69 Although mice are often used as hosts for viral infections, there are no reports of spontaneous viral disease in the eyes or ocular adnexae of mice.

DYSTROPHIC CALCIFICATION This term is used in human ophthalmic pathology to indicate the deposition of mineral salts in tissue either due to elevated levels of serum calcium or associated with areas of cellular injury or necrosis.72 A more appropriate term is dystrophic mineralization, because it is likely that other elements are involved, in addition to calcium. In tissue stained with hematoxylin and eosin (H&E), mineral deposits are densely basophilic and are composed of small or large granules that can form larger aggregates. The von Kossa stain is useful for demonstrating mineralized foci.31 With electron microscopy, the mineral deposits are acellular, irregular, and electron dense. In mice, the cornea and lens are the most common locations for this process. Corneal deposits are generally located in the superficial layers of the corneal stroma (Figure 8.11) and often lead to loss of the overlying corneal epithelium. If this occurs, it is equivalent to a surface defect with all the risks of infection implied (see above). Early corneal mineralization presents clinically as a “smoky” superficial area of haze that is usually 0.5 to 1.0 mm in diameter and is most commonly present in the cornea that is exposed in the palpebral fissure. This location is most likely associated with the greater likelihood of evaporational drying that enhances mineral deposition. Morphologically, the earliest deposits are dustlike and located just below the corneal epithelium. Later, deposits may increase in size to form large aggregates and become clinically larger and more opaque. Stromal scarring near the mineral deposits is a common occurrence. In some mice, focal or diffuse corneal neovascularization may occur.73 Dystrophic mineralization is more often found in mice older than 4 months of age. It has been reported in DBA/2 mice,73,74 but casual strain screening has also identified mineralization in a variety of strains and species, including A/HeJ, BALB/c, CASA/Rk, C3H, LP/J, MOLD/Rk, PERC/Ei, RIII/DmMob, SF/CamEi, WMP/PasDn, and YBR/HeWiHaCvEi (R. Hurd and N. Hawes, personal observations). The variety of affected strains suggests it is likely that mineralization occurs in other strains as well. If located in the visual axis, the corneal opacities can interfere with observation of intraocular structures. Many of these strains also develop cardiac calcinosis and thickened bones (J. P. Sundberg, unpublished observations).

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FIGURE 8.11 Corneal surface disease. A. Clinical photograph of DBA/2J mouse with central superficial corneal mineralization. B. An 11-month-old DBA/2J mouse with superficial basophilic mineral deposits (arrows). Original magnification × 200. C. A.BYSn/J mouse homozygous for the corn1 mutation. At 30 days of age the corneal epithelium is irregularly thickened by coalescent epithelial plaques. Early stromal vascularization is present (arrow). D. By 45 days of age, the cornea is diffusely vascularized. The epithelial changes are still present, but less prominent. E. Typical epithelial plaque in a corn1 mouse. The corneal epithelium is two to three times normal thickness and the most superficial cells are degenerating and loosely adherent to the surface (arrow). Original magnification × 400. F. In an older mouse, superficial neovascularization is a prominent feature (arrow). Original magnification × 400.

CORNEAL NEOVASCULARIZATION Vascular invasion of the normally avascular cornea has been the subject of intensive research for several reasons: 1. Corneal neovascularization is produced by many different stimuli, but understanding of the molecular mechanisms involved remains limited.

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2. Corneal neovascularization is associated with visual loss and failure of corneal grafts. 3. Neovascularization and angiogenesis have pivotal roles in wound healing and in accelerated growth of neoplasms.75 Mice develop both spontaneous and secondary corneal neovascularization and serve as subjects for evaluation of molecular mechanisms of vascular growth. Growth of new vessels into the corneal stroma has been a subject of investigation for many years.76-81 When corneal neovascularization occurs, new vessels originate from the arcade of vessels at the limbus and extend into the cornea, but growth ultimately ceases. Neovascular response to a stimulus and the necessity of its proximity to the limbus for vascular growth to occur76,80 suggest that a diffusible substance may be responsible. A common hypothesis is that there is either upregulation of vasostimulatory substances released by external stimuli or downregulation of vasoinhibitory factors present in normal cornea.82 Although this simple explanation may be correct in principle, the reality is undoubtedly more complex.83 Ease of observation of growing vessels in vivo makes the cornea useful for assessing mechanisms of vascular growth and for testing agents used to inhibit new vessel formation. The most common procedure is to dissect an intrastromal corneal pocket and insert a pellet of ethylene vinyl acetate copolymer (Elvax-40, Alza, Boston, MA) or hydroxyethylmethacrylate (Hydron-Interferon Sciences, New Brunswick, NJ) that contains the material under investigation.84,85 If a substance stimulates angiogenesis, vessels begin to develop at the limbus and extend into the corneal stroma toward the pellet. The extent and degree of vascular growth can be quantified.85 Such techniques have been used to demonstrate the angiogenic activity of basic fibroblast growth factor,86 tumor necrosis factor-α,87,88 TGFα, vascular endothelial growth factor, interleukin-1, and interleukin-8.89 In inflammatory angiogenesis, macrophages that contain many different cytokines play an important role,88,89 as do lymphocytes90 and mast cells.78,91 Clearly, there are a host of factors involved in response to angiogenic stimuli, but the identity, relative importance, and extent of interaction of different factors remain poorly defined. Many human diseases, especially those that affect the corneal surface, provide a stimulus for corneal neovascularization.72,92,93 Disturbances of the corneal surface in mice were described earlier in this chapter. When vascularization occurs, the vessels sometimes remain superficial and lie between the corneal epithelium and stroma. In most cases, however, there is diffuse stromal vascularization, characterized by thin-walled vessels. The vessels may cover the entire cornea or be sectoral, particularly if there is a focal lesion such as a corneal ulcer. Neutrophils, lymphocytes, and plasma cells are typically present during the acute and subacute evolution of the inflammatory process. When the inflammation is related to a foreign body, phagocytic giant cells may be found (see Figure 8.10). Although corneal neovascularization is most often secondary to external factors, it may occur in mice as a result of genetic factors or as a background lesion in some strains. Spontaneous corneal neovascularization has been reported in both nude (Fox1nu) and hairless (hr) mice.94,95 In these mouse mutations, vessels invade the cornea in the perinatal period and progress without inflammation. The vessels persist in adult mice. It has been suggested that these mice demonstrate greater angiogenic activity than normal mice, although the nature of the stimulus is unknown.94,95 An alternative explanation for vascularization in hairless and nude mice is that abnormalities of skin and sebaceous glands (including Meibomian glands) induce the complications of corneal surface disease. Hairless mice develop granulomatous dermatitis, sebaceous gland ductal ectasia, and atrophy of sebaceous glands (including Meibomian glands).1 This creates the potential for corneal surface irritation from desquamating skin flakes (from diffuse hyperkeratosis) that fall onto cornea and conjunctiva. In addition, the loss of sebaceous gland function may interfere with maintenance of the precorneal tear film. Both of these factors could lead to corneal surface damage, increasing the risk of neovascularization. Nude mice also produce excessive flaky debris from hair follicles, enhancing the risk of

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corneal neovascularization.1 Nude mice have structurally weak hair fibers that break and probably contaminate the corneal surface. Although a primary corneal abnormality cannot be excluded, the neovascularization in nude and hairless mice is likely a consequence of the cutaneous and systemic disease characteristic of both strains. Mice homozygous for the autosomal recessive mutation cornea1 (corn1) develop a roughened and opaque corneal epithelium shortly after birth, which is followed by development of corneal neovascularization around 3 weeks of age. The mutated gene is located on mouse Chr 2.56 The mutation occurred in the A.BY/SnJ congenic strain and its effects are limited to the eye. By the second postnatal week, microscopic examination demonstrates mild diffuse thickening of the corneal epithelium that evolves further by the time the eyes open. At P14, there are numerous focal plaques of irregular, cloudy corneal epithelium. Light microscopic examination demonstrates localized thickening and increased eosinophilia of the superficial epithelial cells (Figures 8.11 and 8.12). Cornification (development of a stratum corneum) does not occur. The most superficial epithelial cells lose their mutual cohesion; this is confirmed with scanning electron microscopy. The increase in epithelial thickness reflects an increased rate of cell proliferation that is ten times as high as in heterozygous littermates by 30 days. Stromal neovascularization begins about P18 and reaches the central cornea by P45, when it stabilizes. The vessels are concentrated in the anterior stroma.56 The molecular etiology of the corneal surface disease and neovascularization is under investigation.

KERATOPLASTY Corneal grafts depend on the avascularity of the cornea for maximum visual success, because this shields the transplant from the immune system.96,97 In addition, the cornea exhibits certain forms of immune deviation (reviewed in Streilein et al.98), including the induction of apoptosis of infiltrating cells by FAS-FAS ligand interaction,99 that suppress the immune response. Once this immune privilege is lost, the risk of graft rejection is greatly enhanced.96,100-103 The microscopic appearance of corneal graft rejection in mice is similar to that in other mammals and includes the infiltration of graft and host tissue by mononuclear cells, identified as CD4+ and CD8+ T cells. The CD4+ cells appear to be the principal cell responsible for rejection of mouse corneal grafts.102 The incidence of rejection and the intensity of inflammation are increased when there are anterior synechias attached to the graft margin, presumably because the immune system has better access to the grafted tissue via the iris vessels.103 Corneal endothelial cells are usually destroyed by the inflammatory process, leading to corneal edema. Corneal neovascularization often follows graft rejection and causes additional corneal opacification. In addition, the presence of blood vessels in the rejected graft reduces the success rate for regrafting.

CORNEAL STORAGE DISEASES Diseases in which abnormal proteins, lipids, glycosaminoglycans, and other substances are deposited in the cornea are a common cause of visual disability in humans.104-108 Deposits of abnormal material may be intracellular or extracellular and located in all parts of the cornea, in addition to other ocular and systemic locations. Although less common in mice, two models of human corneal storage disease have been reported. The mucopolysaccharidoses are inherited storage diseases found in many animal species in which lysosomes are filled with an abnormal glycosaminoglycan (reviewed in Sands et al.109). Several chemically different mucopolysaccharidoses have been described in human eyes.31 Mucopolysaccharidosis (MPS) VII occurs in mice and is the biochemical equivalent of the Sly syndrome in humans. MPS VII is caused by β-glucuronidase deficiency. This enzyme breaks down complex glycosaminoglycans and its absence causes them to accumulate within cells of affected animals.109 Mice homozygous for the autosomal recessive mutation (gusmps/gusmps) located on mouse Chr 5

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FIGURE 8.12 Corneal surface disease. A. Unaffected littermate with 2-h exposure to 3H-thymidine to label cells in S-phase. Only a few basal cells of the corneal epithelium (white arrows) are labeled. Original magnification × 250. B. In a homozygous corn1 mouse there is abundant labeling of basal and more superficial cells. Original magnification × 100. C. Unaffected littermate exposed to BrdU shows minimal corneal epithelial labeling. D. A corn1 mouse demonstrating extensive BrdU labeling of the basal corneal epithelium. C and D original magnification × 400. E. View of a corn1 mouse. Relatively normal (N) corneal epithelium is interrupted by a plaque of abnormal epithelial cells that are loosely attached to the surface. F. Scanning electron microscopy of normal A.BYSn/J mouse. The corneal epithelium forms a hexagonal array of thin cohesive cells. An occasional desquamating cell is present (arrow). E and F original magnification × 1800. (E and F, from Smith, R.S. et al., Invest. Ophthalmol. Vis. Sci., 37:397–404, 1996. With permission.)

accumulate intracellular deposits in stromal keratocytes, corneal endothelial cells, and retinal pigment epithelium. Affected cells have a vacuolated appearance (Figure 8.13). Abnormal lysosomal deposits are also present in the central nervous system, spleen, liver, and bone (osteoblasts, osteocytes, chondrocytes, and bone marrow cells).109 Affected mice have shortened life spans, dwarfism, decreased joint mobility, and extensive skeletal deformation.110 Gene therapy with intravitreal injection

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FIGURE 8.13 MPS VII mice (gusmps/gusmps). A. The cytoplasm of the nonpigmented ciliary epithelium is distended with deposits (arrow) of the specific glycosaminoglycan. B and C. The corneal endothelium (arrow-B) and stroma (arrow-C) contain similar deposits. A to C original magnification × 200. D. The retina is normal, but the retinal pigmented epithelium contains glycosaminoglycan deposits (arrow). Original magnification × 400. E. An 8-month-old mouse on 1% cholestanol diet. There is a central granular corneal opacity (*), toward which new vessels are extending (arrows). Original magnification × 240. F. Directly beneath the corneal epithelium (Ep) there is a rectangular plate-shaped array of crystals (inset arrows). New vessels (v) are present in the corneal stroma (St) of a cholestanol-treated mouse. Original magnification × 600. (A to D, courtesy of Dr. Carole Vogler. E and F, courtesy of Dr. Yousuke Seyama; reprinted from Kim, K.S. et al., Biochim. Biophys. Acta, 1085:343–349. © 1991. With permission of Elsevier Science.)

of recombinant adenovirus111 and bone marrow transplantation are partially effective forms of treatment for the ocular component of this disease in mice, as is early enzyme replacement treatment with β-glucuronidase.112

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Other exogenous material may also be deposited in the cornea. An interesting model of a corneal dystrophy resulting from cholestanol (the 5-dihydro derivative of cholesterol) feeding of mice113 is a phenocopy of the human disease known as Schnyder’s crystalline corneal dystrophy (SCCD).106,114 SCCD is an autosomal dominant, bilateral corneal disease characterized by corneal clouding, arcus lipoides (circumferential stromal lipid deposits located at the limbus), and crystalline cholesterol deposits located in the superficial corneal stroma. The gene for SCCD maps to human chromosome 1p34.1–p36.106 BALB/cJ mice fed a cholestanol-enriched diet for 8 months developed amorphous conglomerates of calcium and phosphate as well as rectangular cholestanol crystals beneath the corneal epithelium (see Figure 8.13).113 In clinical appearance these crystals have a strong resemblance to those found in SCCD that are known to be composed of a mixture of phospholipid and cholesterol compounds.106

CORNEAL EDEMA Corneal opacification results from any process that disturbs epithelial integrity or the regular arrangement of stromal lamellae. In addition to the causes of corneal cloudiness already discussed, generalized stromal edema interferes with both vision and visualization of structures within the eye. In most cases, functional failure of the corneal endothelium initiates development of corneal edema. The monolayer of corneal endothelium pumps fluid and electrolytes out of the stroma and maintains a barrier between the stroma and the anterior chamber. The end result is lower water content of the corneal stroma compared to other tissues. Even small shifts in corneal fluid balance alter the regular arrangement of stromal lamellae and cause corneal cloudiness. As a consequence, both epithelial and stromal edema may develop. In human eyes, corneal edema from endothelial decompensation is a major cause of visual dysfunction. Often, the decompensation is spontaneous or occurs postoperatively in individuals with Fuchs’ endothelial dystrophy. Because the human corneal endothelium does not undergo mitosis, the normal decrease that occurs with aging places endothelial function at greater risk. In Fuchs’ dystrophy, there is both a decrease in total endothelial cell count and a disturbance in the regular hexagonal array of endothelium.72,96,115,116 Both of these findings are associated with deficient endothelial function. A second characteristic of Fuchs’ dystrophy is the occurrence of many areas of focal Descemet’s membrane thickening (cornea guttata).96 Corneal endothelial decompensation can develop following trauma or be induced by severe inflammation as occurs in acute corneal graft rejection (see above). Mice and rats demonstrate progressive loss of corneal endothelial cells and loss of the normal hexagonal form with advancing age. Although corneal edema related to endothelial cell functional loss does not develop in either mice or rats, cell loss between 1 and 30 months of age has been reported to be as much as 36% in C57BL/6J mice.117 Patchy corneal edema, likely related to trauma, is infrequent (Figure 8.14). Focal Descemet’s membrane thickening occurs in mice (see Figure 8.14) as an isolated event, rather than multiple deposits as found in Fuchs’ dystrophy in human eyes. When corneal stromal edema occurs in mice it is usually a developmental defect, rather than agerelated. A transgenic mouse line that carries a rearranged T-cell receptor δ transgene clinically mimics human corneal decompensation. As early as E18.5 abnormally thin corneal epithelium is demonstrated. By P8, peripheral corneal edema develops and by P21 there is generalized corneal epithelial edema. Adult mice demonstrate continued intercellular epithelial edema, stromal edema, and marked endothelial cell loss.118 There are several mechanisms that might be evoked in this model, but the pivotal feature is the major loss of corneal endothelial cells that control corneal hydration; the other changes represent secondary effects of endothelial dysfunction. Development of the corneal stroma and endothelium depends on migration and differentiation of neural crest119 and cranial paraxial mesoderm120 during embryogenesis. It is likely that these processes are regulated by multiple genes, alterations of which lead to a spectrum of congenital abnormalities.

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FIGURE 8.14 Corneal edema/endothelialization. A. Peripheral corneal edema in C57BLKS/J mouse, possibly of traumatic origin. B. The anterior two thirds of the corneal stroma is edematous (*), as indicated by the decreased eosin staining. Epithelial edema is absent and the posterior stroma is uninvolved. Original magnification × 400. C. In chronic anterior corneal stromal edema, neovascularization may develop (arrow). Original magnification × 400. D. An isolated area of focal proliferation of Descemet’s membrane in a C57BL/6J mouse (arrow). Original magnification × 630. E. In a DBA/2J mouse Descemet’s membrane has proliferated over the anterior iris surface, causing an anterior synechia (arrow). Original magnification × 400. F. A similar finding in an AKXD-28/Ty mouse. The arrow indicates a corneal endothelial cell on the new-formed Descemet’s membrane extending onto the iris surface. Original magnification × 630.

For example, null mutations of the mouse gene, Foxc1 (forkhead box C1) result in total failure of formation of the corneal endothelium, thickening and irregular collagen deposition in the corneal stroma, and increased thickness of the corneal epithelium with intercellular edema (Figures 8.15 through 8.17).121 Similar ocular changes were identified in transgenic mice that expressed either TGFα or EGF in the lens.122 In all cases of corneal edema the common denominator is diminished or absent endothelial pumping function, whether related to trauma, development, or age.

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FIGURE 8.15 Embryos heterozygous and homozygous for congenital hydrocephalus (Foxc1ch, formerly Mf1ch) at E17. A. The eyelids (EL) of the ch/+ mouse are closed and there is a well-formed anterior chamber (*). Abbreviations: cornea (C), trabecular meshwork (TM), lens (L), retina (R), sclera (S). B. In the homozygous mutant, the lids are open and the anterior chamber is absent. A and B original magnification × 20. C. Enlargement of boxed area demonstrates a distinct endothelial layer (arrow). D. Both the corneal endothelium and the anterior chamber are absent in the homozygous mutant. C and D original magnification × 400. (From Kidson, S.H. et al., Dev. Biol., 211:306–322, 1999. With permission of Academic Press.)

CORNEAL ENDOTHELIAL PROLIFERATION Although the corneal endothelium usually responds to stimuli by diminished function and cell death, on occasion, corneal endothelial cells proliferate and migrate over the surface of the trabecular meshwork and iris. When these displaced endothelial cells produce thickened Descemet’s membrane-like material, the phenomenon is known as endothelialization, which in human eyes is associated with some forms of glaucoma, trauma, and with the iridocorneal endothelial syndrome.72,123,124 Endothelialization of the anterior chamber angle occurs in mice of the DBA/2J and AKXD-28/Ty strains, both of which develop progressive angle closure glaucoma (Figures 8.14 and 8.18).74,125

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FIGURE 8.16 Heterozygous and homozygous congenital hydrocephalus (FoxC1ch). A and B. At E13.5 normal and mutant cornea are composed of a thin corneal epithelium (EP) and mesenchyme (CM). Small anterior hyaloid vessels (arrow) abut the lens. In A, there is evidence of early anterior chamber formation (*). C and D. At E17.5, the normal cornea has separated from the lens and there is a well-defined endothelial layer (EN). D. The epithelium of the ch/ch mouse is thickened and has an irregular surface. The cornea of the mutant is still attached to the lens and corneal endothelium is absent. E. At E18.5 the cornea of the +/+ mouse is developing normally. The corneal stroma is arranged in parallel arrays. F. The ch/ch mouse shows persistent epithelial thickening and there are many large intercellular spaces. The corneal stroma is disorganized and the keratocytes have a stellate appearance. Blood vessels are present in the corneal stroma (arrowhead). Original magnification of all pictures × 400. (From Kidson, S.H. et al., Dev. Biol., 211:306–322, 1999. With permission of Academic Press.)

AQUEOUS OUTFLOW AND INTRAOCULAR PRESSURE The mouse eye maintains a steady range of intraocular pressure that varies from strain to strain,126 usually between 10 and 20 mmHg. This homeostatic system depends on secretion of aqueous humor by the ciliary body, its unimpeded flow into the anterior chamber, and its outflow from the eye through the trabecular meshwork, Schlemm’s canal,127 and the uveoscleral pathway.128 Anything that

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FIGURE 8.17 Null mutation of the transcription factor Foxc1. Corneal epithelium and stroma at E18.5. A. Corneal epithelium of the mutant mouse is five cells thick. The superficial cells are enlarged and demonstrate swollen intercellular spaces (*). Bowman’s layer is absent. B. The corneal epithelium of a normal littermate is two to three cells thick and rests on a well-formed Bowman’s layer (arrow). A and B original magnification × 4000. C. The corneal stroma of a normal mouse demonstrates flattened keratocytes arranged parallel to the surface and separated by bundles of collagen fibers arranged at right angles to each other. D. Keratocytes of mutant mice are rounded or stellate, disorganized, and separated by irregular swirls of collagen. C and D original magnification × 7000. (From Kidson, S.H. et al., Dev. Biol., 211:306–322, 1999. With permission of Academic Press.)

interferes with aqueous outflow leads to elevated intraocular pressure, which is frequently associated with glaucoma. Glaucoma has widespread ocular effects and its signature is optic nerve damage, mediated in part by retinal ganglion cell death. Glaucoma is a complex disease in both mice and humans with manifold molecular mechanisms and multiple risk factors101,129 (glaucoma in mice is reviewed in John et al.130).

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FIGURE 8.18 A. Causes of glaucoma. A. Anterior synechias may develop as glaucoma progresses, resulting in a “false” anterior chamber angle anterior to the true location of the trabecular meshwork (arrow). All evidence of Schlemm’s canal and the trabecular meshwork have been obliterated. B. Although Schlemm’s canal and the trabecular meshwork are visible (arrow), a broad anterior synechia renders them nonfunctional. A and B original magnification × 200. C. Posterior synechias have firmly attached the iris to the lens in this AKXD-28/Ty mouse (arrows). An anterior subcapsular cataract is present. D. If the posterior synechias completely surround the pupil, aqueous will be trapped in the posterior chamber, causing the iris to balloon forward—an iris bombe. An eosinophilic exudate fills the anterior chamber. The iris leaflets are indicated by arrows. C and D original magnification × 100. E. In a 26-month-old DBA/2J mouse, access to the trabecular meshwork is blocked by an anterior synechia associated with extension of Descemet’s membrane (arrow) across the angle structures and onto the anterior iris surface. Original magnification × 400. F. In a 9.5-month-old DBA/2J mouse, Descemet’s membrane (arrow) and the corneal endothelium cover the posterior corneal (C) surface. An endothelial cell (E) lies on the iris surface and newly formed Descemet’s membrane-like material (arrowhead) continues across the iris surface. Original magnification × 8100.

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CLASSIFICATION OF GLAUCOMA There are numerous possible classifications of glaucoma;101 a reasonable subdivision based on clinical evaluation and histopathology is developmental, open angle, and closed angle. For those unfamiliar with the term angle, it refers to the actual angle formed by the anterior surface of the iris and the posterior surface of the cornea. Schlemm’s canal and the trabecular meshwork are located within the corneoscleral junction external to the angle recess, the aqueous veins are external to Schlemm’s canal, and the bulk of the ciliary muscle and ciliary processes lie behind the angle (see Chapter 1). Developmental Glaucoma As described in Chapter 3, development of the trabecular meshwork and Schlemm’s canal is an intricate process involving interaction between the peripheral edge of the optic cup, mesenchyme derived from neural crest, and cranial paraxial mesoderm. Development of the outflow structures is nearly complete at birth in human eyes, although minor remodeling may continue for 2 to 3 years after birth.131,132 In mice and other animals important developmental changes continue for several months after birth.133 Developmental remodeling of the trabecular meshwork and Schlemm’s canal in mice is not complete until 5 to 8 weeks after birth.134 Considering the intricacies of development of the outflow channels, developmental abnormalities of these structures are not surprising. The developmental glaucomas are a group of diseases associated with abnormalities in the structure of the angle components. Elevated intraocular pressure and nerve damage are variably associated with typical morphological changes. Absence of Schlemm’s canal, trabecular meshwork hypoplasia, and aberrant tissue deposition have all been described in mice (see below).135,136 Open Angle Glaucoma The most common form of human glaucoma is primary open angle glaucoma and it is one of the principal worldwide causes of blindness.137 Although knowledge is advancing rapidly, glaucoma is a complex disease with many poorly understood risk factors and no clearly identified molecular mechanisms.130,138 It is known that mean intraocular pressure levels differ in different mouse strains,126,130 and this suggests that selective breeding or techniques of genetic engineering may be capable of producing a model of primary open angle glaucoma. Such a model does not currently exist. Angle Closure Glaucoma In human and mouse eyes, glaucoma may develop when the anterior chamber angle (formed by the angle between iris and cornea) closes, covering the trabecular meshwork and Schlemm’s canal. When most of the angle becomes closed (iris and cornea in contact and often adherent to each other), then aqueous can no longer reach the trabecular meshwork and elevated intraocular pressure develops. This may occur because of anatomically narrow angles (primary angle closure glaucoma) or because of a process that produces anterior or posterior synechias (iridocorneal or iridolenticular adhesions) that obstruct access to the chamber angle (secondary angle closure glaucoma).72 Although mice have relatively narrow anterior chamber angles, current technology does not allow detection of primary angle closure glaucoma. Secondary angle closure occurs in several inbred strains, some of which develop elevated intraocular pressure and sequelae similar to those reported in human glaucoma.74,139

ANTERIOR SEGMENT FINDINGS IN GLAUCOMA Many parts of the eye are affected by glaucoma, although the extent of involvement varies with the type of glaucoma as well as its duration. Most anterior segment lesions reported in human glaucoma72

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have been reported in mice with glaucoma74,139 or that carry mutations associated with human glaucoma.135,136 The specific mouse models are discussed later in this chapter. Adhesion of the iris to other ocular structures is commonly associated with secondary angle closure glaucoma, although it also occurs after inflammation or trauma. Iris adhesions are referred to as synechias. As the process begins, the peripheral iris adheres to the overlying cornea because of its proximity, forming peripheral anterior synechias. Anterior synechias may be focal or may eventually attach the entire iris to the posterior corneal surface (see Figure 8.18). When most of the angle is occluded by synechias, there is a strong risk of increased intraocular pressure. A second consequence of synechia formation is destruction of the corneal endothelium in the area of adhesion. The loss of corneal endothelium interferes with corneal fluid balance and may result in corneal edema. Although this has been observed, it is uncommon in mice, perhaps because the unaffected portions of corneal endothelium are able to keep the affected cornea relatively dehydrated. Additionally, with partial synechias, there is likely sufficient function to maintain stable corneal hydration. Corneal and epithelial edema may also occur with acute elevation of intraocular pressure. In some instances, only small anterior synechias form and for unknown reasons, the corneal endothelium is stimulated to migrate across the surface of the trabecular meshwork and peripheral iris. The migrating endothelium retains its capacity to produce basal lamina and Descemet’s membrane-like material (see Figure 8.18). This is referred to as endothelialization and its occurrence effectively blocks the trabecular meshwork. The iris may also adhere to the lens, forming posterior synechias. In most cases, posterior synechias are focal, but they may involve the complete circumference of the pupillary border, which will block access of the aqueous humor to the anterior chamber. Since aqueous production continues, the iris may balloon forward and intraocular pressure will increase (pupillary block glaucoma). This may be so extreme that the anterior iris surface occludes the trabecular meshwork. When complete posterior synechias are present, fibrovascular tissue arising from the iris often extends across the pupil, forming a pupillary membrane (see Figure 8.18). Normal drainage of aqueous humor is hampered by deposition of cells or particulate matter in the trabecular meshwork that may cause additional damage by reaction of trabecular meshwork tissue to the alien material. Macrophages filled with debris, tumor cells, and particulate matter may cause elevated intraocular pressure in human eyes.72 The trabecular meshwork may be plugged by erythrocytes in a hyphema (anterior chamber hemorrhage) or as a secondary result of ingestion of red cell debris by macrophages that accumulate in the trabecular meshwork (Figure 8.19). In similar fashion, following a vitreous hemorrhage, debris-filled macrophages may flow forward with the aqueous and lodge in the trabecular meshwork. The end result of any of these processes may be secondary open angle glaucoma or secondary closed angle glaucoma if synechias develop. The most commonly observed material obstructing the trabecular meshwork is intra- or extracellular melanin pigment. Both the iris pigment epithelium and iris stromal melanocytes may be damaged by disease and die, liberating pigment granules and cell debris that are subsequently ingested by phagocytes. It is normal to identify small numbers of phagocytic cells in the iris stroma and a few are found in or close to the trabecular meshwork of normal mice. In normal mice, cells containing phagocytosed pigment are most commonly located near the iris sphincter or near the iris root where it attaches to the sclera. Cells with phagocytosed melanosomes are typical of mice homozygous for the iris pigment dispersion (ipd) or iris stromal atrophy (isa) loci that are associated with glaucoma in aging DBA/2J mice (see Figure 8.19).139 Late in the disease, these cells sometimes accumulate in such numbers that in addition to the trabecular meshwork, the phagocytes invade the suprachoroid space and sclera over the peripheral retina. The endothelial cells of the trabecular meshwork are capable of phagocytosing pigment granules, although development of glaucoma could not be directly related to mechanical effects of pigment accumulation.140 Ingestion of free melanin pigment by trabecular endothelial cells might alter their metabolism and indirectly interfere with function. Another possible explanation is that toxic intermediates in melanin synthesis could damage the trabecular

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FIGURE 8.19 Secondary glaucoma. A. In a mouse homozygous for the gene for the pigment dispersion gene (ipd), the anterior chamber angle contains many cells (presumably macrophages) distended with pigment. B. These pigment-laden cells may collect in the depths of the angle and occlude the trabecular meshwork, causing secondary glaucoma (methylene blue stain). A and B original magnification × 400. C. The anterior chamber may become filled with blood (*) either spontaneously as in this mouse or subsequent to trauma. Original magnification × 200. D. The densely packed blood (*) may occlude outflow through the trabecular meshwork, causing a secondary glaucoma. The lens (L) is normal. Original magnification × 400.

meshwork endothelium and other ocular structures by mechanisms of cell death, as occurs in hair follicle melanocytes of mice carrying the light allele of tyrosinase related protein (Tyrp1B-lt).130,141 The pigment-filled cells and free melanin pigment in the angle arise either from the pigmented stromal melanocytes, from the posterior iris pigment epithelium, or from macrophages. The observation that multiple melanosomes are often found contained within a unit membrane structure supports the hypothesis that pigment has been ingested by the cell, but does not identify the histogenesis of the cell. Changes in both stromal melanocytes and iris pigment epithelial cells occur in aging DBA/2J mice and their derivative strains,74,139 as well as in human eyes with pigmentary glaucoma.100 The iris stroma becomes diffusely atrophic as a secondary change in chronic glaucoma. Extracellular matrix, vascular channels, sphincter and dilator muscles, and stromal melanocytes are all affected, gradually becoming atrophic to a point where the stroma is barely recognizable. The molecular mechanisms of the underlying processes are poorly understood. A period of prolonged elevation of intraocular pressure may be followed by a return to normotensive or even hypotensive readings.142,143 The reason for the pressure drop is progressive atrophy of the ciliary body that results in decrease or cessation of aqueous production. A similar explanation (see Figure 8.22) likely accounts for the pressure drop with increasing age in DBA/2J mice.74 Care must be taken in making this morphological diagnosis because appropriate tissue orientation is critical for accuracy. Review of multiple step sections is necessary before a conclusion can be reached (see Chapter 13).136 The first obvious sign of atrophy is flattening of the ciliary processes followed by

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shrinkage of the connective tissue and vasculature beneath the ciliary epithelium. In extreme cases, only a small tag of epithelial tissue remains.

POSTERIOR SEGMENT FINDINGS IN GLAUCOMA RETINA AND OPTIC NERVE Although monitoring of intraocular pressure and its normalization were central tenets of glaucoma treatment for many years, much attention is now focused on events occurring in the retina and optic nerve. Distribution of retinal ganglion cells is not even across the retina. Near the optic nerve, the ganglion cells are closely spaced, forming a layer two to four cells thick. A short distance from the optic nerve, this becomes a single layer of cells that remain closely packed. Anterior to the equator, the nuclei become more widely spaced (Figure 8.20). Therefore, it is important when assessing the health of the retinal ganglion cell layer in mice to do so in a consistent fashion to avoid interpretation errors. Ideally, the sections evaluated should pass through the center of the cornea and through the optic nerve to avoid obliquity (see Chapter 13).136 In most types of glaucoma, retinal ganglion cell loss is progressive and in severe glaucoma, most cells disappear. Since the ganglion cells give rise to the axons of the nerve fiber layer, there is concomitant thinning of this layer as well (see Figure 8.20). Death of retinal ganglion cells and loss of the nerve fiber layer are associated with atrophic changes in the optic nerve. Fundus photography is a useful technique to follow the development of optic nerve cupping (see Figure 8.20). There is normally a shallow cup on the surface of the mouse optic nerve, which does not extend much below the level of the ganglion cell layer. In advanced cupping, the excavation deepens posteriorly to a level parallel to the photoreceptor layer. The mouse lamina cribrosa is more delicate than that of the primate, but is definitely present (see Chapter 2). Posterior bowing of the lamina cribrosa can be identified in advanced cupping. Posterior to the lamina cribrosa, chronic glaucoma is associated with collapse and atrophy of the nerve fiber columns and irregularity of the pial septae. Atrophy of the nerve may cause a decrease in nerve diameter to 25% of normal.74 Special stains such as paraphenylenediamine144-146 demonstrate more striking early axonal loss than would be suspected with routine H&E staining. In advanced stages of glaucoma, few viable axons remain (see Figure 8.20). Optic atrophy is frequently accompanied by glial proliferation in both pre- and postlaminar optic nerve. This may become so extensive that it obscures the cupped optic nerve.

GLAUCOMA AND GLAUCOMA-LIKE EFFECTS IN MICE Neither physical nor genetic factors predisposing to glaucoma necessarily produce elevated intraocular pressure and glaucomatous neuropathy. For example, the glaucoma that frequently occurs following blunt injury to the eye does not always develop, despite seemingly severe injury.147,148 The explanation lies in the extent of traumatic injury to anterior chamber angle structures. For example, there may be extensive trabecular meshwork damage in one area, but elsewhere there is enough uninjured angle to permit maintenance of normal intraocular pressure. A similar disparity is seen when a patient with a known mutation associated with glaucoma never develops elevated intraocular pressure or glaucomatous optic neuropathy.149,150 A potential explanation similar to that for traumatic glaucoma has been suggested in mice heterozygous for the forkhead transcription factor genes Foxc1 and Foxc2.93 The homologous human gene, FOXC1, is associated with some forms of Axenfeld–Rieger syndrome (ARA).117 Foxc1 and Foxc2 mutant mice do not develop elevated intraocular pressure on the genetic backgrounds studied, likely due to the presence of both normal and abnormal stretches of trabecular meshwork and iridocorneal angle in each eye. Similarly, not all patients with ARA develop elevated pressure and optic neuropathy. In addition, genetic background, as discussed in Chapter 6,

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FIGURE 8.20 Effects of glaucoma. A. In a normal retina, near the optic nerve, retinal ganglion cells are tightly packed in a one to two cell thick layer (arrow). The inner nuclear layer is five to six cells thick. B. In an older mouse homozygous for the Tyrp1b allele, with chronic glaucoma, ganglion cells are sparse (arrow). Nearly half of the inner nuclear layer (arrowhead) has also been lost. A and B original magnification × 400. C. In a normal optic nerve, there is a shallow depression associated with the retinal vessels (V) that ends at the level of the ganglion cell layer (arrows). D. In an AKXD28/Ty mouse with longstanding glaucoma, there is advanced cupping (arrowheads) that extends posteriorly behind the depth demarcated by the choroid (C). (C and D, courtesy of Michael Anderson and BioMed Central, 2:1–12, 2001.) Original magnification × 200. E. In a normal optic nerve from a 2-month-old DBA/2J mouse there is modest variation in axon size, but all axons are surrounded by a sharply defined ring of myelin and the axons are arranged in regular bundles. F. In an older, severely affected DBA/2J mouse the optic nerve is completely disorganized and the majority of axons stain darkly with paraphenylenediamine (see Chapter 13), indicating advanced degeneration (arrows). Original magnification × 630. (E and F, courtesy of Abbot Clark, Alcon Laboratories.) G. Normal fundus photograph. H. Glaucomatous fundus with an asymmetric and severely cupped optic nerve (arrowhead) as well as peripapillary chorioretinal atrophy. (G and H, courtesy of Michael Anderson and BioMed Central, 2:1–12, 2001.)

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exerts a major effect on the phenotypic consequences of the mutation. For these reasons, we will discuss not only glaucoma in mice, but also morphological changes frequently associated with glaucoma that are found in several inbred strains.

AGE-RELATED GLAUCOMA IN DBA/2J AND RELATED STRAINS In young DBA/2J (D2) mice, the iris and other ocular structures appear normal on slit lamp examination. By 4 months, peripheral transillumination is visible and becomes more prominent with progression of time (Figure 8.21). The clinical appearance is due to progressive atrophy of the iris stroma and epithelium, including pigment dispersion. Large cells filled with pigment accumulate in the deep angle recess concomitant with development of anterior synechias. In some mice, endothelialization of the anterior chamber angle covers the synechias (see Figure 8.18). There may be prominent accumulations of pigment-laden cells adjacent to the iris sphincter (Figure 8.22). At the same time there is ongoing atrophy of the ciliary body, which nearly disappears in older mice.74 The anterior segment of D2 mice often appears enlarged on slit lamp examination, a finding confirmed by ophthalmic ultrasound.71 The high frequency of central corneal mineralization often impedes fundus examination. Nuclear and anterior subcapsular cataracts are found in older D2 mice. In addition, some of the oldest mice demonstrate subluxation of the lens, usually when there is extreme

FIGURE 8.21 Clinical phenotype changes with age and genetic background. A to C. AKXD 28/Ty mice; D to F. DBA/2J mice. The AKXD-28/Ty exhibit only the iris stromal atrophy (isa) phenotype, whereas the DBA/2J mice show both isa and iris pigment dispersion (ipd) phenotypes. A and D. Two-month-old mice of both strains are clinically normal. B and E. By 12 months of age, AKXD 28/Ty mice (B) have iris stromal atrophy with loss of iris detail, exposure of the sphincter muscle (arrowhead), and early transillumination defects (arrow). DBA/2J (E) mice of similar age have both iris stromal atrophy and severe pigment dispersion as well as posterior synechias and pigment deposits on the lens and iris (arrowheads). C and F. In 26-month-old mice of both strains there is severe iris stromal atrophy with irregular pupils and iris holes. Although the AKXD 28/Ty iris transilluminates, the effect is much greater in the DBA/2J iris because of the combined loss of stromal and epithelial pigment combined with severe stromal atrophy. (Courtesy of Michael Anderson and BioMed Central, 2:1–12, 2001.)

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FIGURE 8.22 A. Glaucoma in DBA/2J mice. In a young mouse, the optic nerve remains normal. The peripapillary nerve fiber layer is evident entering the optic nerve (arrows) and the axons in the nerve are arranged in regular columns by the pial septae (arrowheads). Original magnification × 100. B. In a 22-month-old mouse, the retrobulbar optic nerve is atrophic and disorganized. Taking the magnification into account, this optic nerve is less than 25% the diameter of the healthy nerve (A). Original magnification × 200. C. The iris of a 3-month-old mouse shows a robust stroma, pigment epithelium, and iris. D. In the same 3-month-old mouse, Schlemm’s canal (arrow), trabecular meshwork, and ciliary body are normal. C to D original magnification × 200. E. In an 8-month-old mouse there is a small anterior synechia. The iris pigment epithelium is quite atrophic, and in some locations (arrowhead) there is marked thinning of both stroma and PE. Schlemm’s canal (arrow) is shorter than normal. Original magnification × 100. F. In a 20-month-old mouse, a broad anterior synechia is present. Both iris stroma and pigment epithelium are atrophic. Original magnification × 100. G. Extensive accumulations of pigment-laden cells have produced large nodules in the iris sphincter (arrow) Original magnification × 100. H. In a 20-month-old mouse the ciliary body (arrowhead) is atrophic and the zonules (arrow) lack normal tension. This latter finding reflected the presence of a lens dislocation. Original magnification × 200.

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ciliary body atrophy and zonular relaxation. Ciliary body atrophy accompanied by decrease in aqueous humor production is the likely explanation for the fall of intraocular pressure to subnormal levels that occurs in older mice (see Figure 8.22).74 Not long after elevated intraocular pressure is detected between 6 and 9 months of age, the retinal ganglion cells and nerve fiber layer begin to disappear. At the same time, the delicate structure of the lamina cribrosa and orderly array of retrolaminar nerve fiber bundles is lost. Optic nerve atrophy and cupping are first identified at 11 months and become more common in older mice. In some cases, the cupped optic nerve is obscured by gliosis. The changes in the optic nerve are reflected in cross sections of the nerve in older mice, which show progressive axonal damage (see Figures 8.20 and 8.22).74 The clinical and morphological phenotype in DBA/2J mice includes features that resemble pigment dispersion syndrome151 as well as the iridocorneal endothelial syndrome152 in human eyes. When DBA/2J (D2) mice are crossed with C57BL/6J (B6) mice and their progeny either intercrossed or backcrossed to DBA/2J, mice are produced that contain either D2 or B6 alleles or a combination of the two. The backcross mice segregate into four different clinical phenotypes: normal, pigment dispersion, iris atrophy, and a combination of both that resemble the original D2 phenotype. Two loci were identified, one on mouse Chr 4 (named isa for iris stromal atrophy) and the other on Chr 6 (named ipd for iris pigment dispersion). The clinical phenotypes correspond to the genotypes: normal mice are heterozygous at both loci; pigment dispersion mice are homozygous for the ipd allele of D2; mice with iris atrophy are homozygous for the D2 isa allele, and the most severely affected mice are homozygous for both D2 alleles.139 The morphologic phenotype also corresponds to the genotypes. The ipd phenotype demonstrates substantial loss of the iris pigment epithelium. The isa phenotype has little remaining iris stroma but good preservation of iris pigment epithelium. The most severely affected mice, doubly homozygous for D2 alleles, resemble the original DBA/2J mice (Figures 8.21 through 8.23). Retinal and optic nerve changes are similar to those described in DBA/2J mice. Discovery that the tyrosinase-related protein gene (Tyrp1) is a candidate gene for isa has helped detect other mouse strains with anterior segment abnormalities similar to those of D2. The Tyrp1b allele of D2 mice is associated with a brown coat color and is always associated with iris stromal atrophy. A number of inbred brown strains also demonstrate this phenotype, including C57L/J, DBA/1J, SEC/1ReJ, and all brown coat color BXD (B6xD2) recombinant inbred strains. The iris atrophy phenotype is absent in all BXD strains with black coat color, which lack the Tyrp1b allele.97 Pigmented BALB/cByJ (pigmented BALB) congenic for the wild-type tyrosinase allele and carrying the Tyrp1b allele characteristic of BALB/c also develop iris stromal atrophy and iris holes, whereas ordinary albino BALB/cByJ do not, demonstrating the importance of pigment production (see Figure 8.22) to the phenotype.139 The recombinant inbred (RI) strain AKXD-28/Ty was generated from an initial cross of mice of the AKR/J and D2 strains.153 The genome of these mice is derived from both parental strains, and the RI strain is homozygous for D2 alleles at many loci including those for isa and ipd. In many ways, the clinical and morphological phenotype is similar to that described in D2 mice, except that free pigment on the anterior lens capsule, a feature typically seen in D2 mice, is missing. Extensive anterior synechias, endothelialization of the angle, and pigment accumulation in the trabecular meshwork occur. Based on the preservation of the iris pigment epithelium, the angle pigment is likely derived from iris stromal melanocytes and released as the stroma undergoes severe atrophy. Retinal damage is more severe than in D2 mice and the outer retina is affected in older mice, with loss of both inner and outer nuclear layer cells. Extensive cupping of the optic nerve is more common and more severe than in D2 mice. Since AKXD-28/Ty mice carry chromosomal markers associated with both ipd and isa, it is curious that they fail to develop the ipd phenotype that displays massive peripupillary

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FIGURE 8.23 Iris phenotypes in the isa and ipd mutants. A. In mice heterozygous at both loci, the iris is normal with robust stroma and pigment epithelium. B. In mice homozygous for the ipd mutation the iris stroma is robust, but contains many pigment-laden cells. The almost total loss of the iris sphincter (arrow) that is normally located between the stroma and pigment epithelium emphasizes the absence of pigment epithelium. C. In mice homozygous for the isa mutation, the pigment epithelium is well maintained, but the iris stroma (arrow) is atrophic and contains only a few pigment-filled cells. D. In mice homozygous for both loci, both iris stroma and pigment epithelium are thinner than normal. The importance of pigment production for the isa phenotype is demonstrated by the normal appearance of the iris in a BALB/cByJ mouse that carries the candidate gene for the isa locus, Tyrp1b, on an albino background. E. In an unpigmented BALB mouse 24 months of age, the iris and angle structures are completely normal. F. A pigmented BALB mouse of similar age has developed iris holes (arrow) and extreme iris atrophy. A to H original magnification × 400.

accumulations of pigment-filled cells. These observations suggest the presence of one or more modifier genes (see Figure 8.21).125

FORKHEAD TRANSCRIPTION FACTORS AND GLAUCOMA In human patients, the ARA anomaly includes developmental abnormalities of the anterior chamber with variable ocular morphological phenotypes and is often associated with glaucoma.154-156 Mutations in the human forkhead gene FOXC1 have been described in patients with several glaucoma phenotypes.157,158 Haploinsufficiency of mouse forkhead genes Foxc1 (forkhead box C1—the mouse homologue of human FOXC1) and Foxc2 produce morphological phenotypes that resemble ARA anomaly.135 Congenital hydrocephalus (ch) in mice is due to a truncating mutation of Foxc1.159,160 Skeletal, cardiovascular, and ocular abnormalities occur in addition to hydrocephalus in Foxc1ch/ch and Foxc1 null mutants. The eyelids of these mice are open at birth resulting in corneal exposure. Foxc1–/– mice fail to develop an anterior chamber and the corneal endothelium is lacking. The absence of

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corneal endothelium is likely due to a failure of migration of neural crest cells responsible for its development. Corneal stromal collagen is irregularly arranged and the corneal epithelium is thickened, in part by intracellular edema (Figures 8.15 through 8.17).121 Because Foxc1–/–, Foxc1ch/ch , and Foxc2 mutant mice usually die in the pre- or perinatal period the full clinical phenotype cannot develop. Heterozygous Foxc1+/–, Foxc1ch/+, and Foxc2+/– survive and form more complex clinical and morphological phenotypes. The ocular clinical phenotypes are limited to the anterior segment of the eye (Figure 8.24). The pupils of affected mice are often irregular and peripherally displaced. Dense iris processes attach to the peripheral cornea that is often opaque (scleralized) (Figures 8.25 and 8.26).

FIGURE 8.24 Clinical phenotypes of Foxc1 mutant mice. Clinical phenotypes are shown for mice of the indicated genetic backgrounds and genotypes (B6-C57BL/6J; 129-129/SvEvTac; CH-CHMU/Le). A and B. The eyes are normal with small, round central pupils. There is a small inferior notch in the B6 pupil, as sometimes occurs in wild-type mice of this strain. C. The pupil in this heterozygote is drawn to one side, where Schwalbe’s line is misplaced (anterior embryotoxon, arrow). D. The pupil is elongated and the iris pigment epithelium is displaced (ectropion uveae, asterisks). This may be caused by two areas of peripheral iridocorneal attachment (arrows) that appear to exert traction on the iris. E and F. In both heterozygotes, the pupil is irregularly shaped and displaced. Both eyes show scleralization of the cornea that is more prominent in F (arrow). Also in F, there is an irregular corneal opacity (arrowheads) due to attachment of the inferior iris to the posterior corneal surface. (From Smith, R., Hum. Mol. Genet., 9:1021–1032, 2000. With permission of Oxford University Press.)

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FIGURE 8.25 Iridocorneal angle defects in Foxc1 mutant mice. The strain abbreviations are the same as in Figure 8.24, with the addition of 129BS, a strain of mixed 129 and Black Swiss background. A and B. In wildtype mice, Schlemm’s canal (SC) extends from a point above the posterior ciliary body forward to a point close to the cornea (arrowheads). The trabecular meshwork (TM; arrows) is normal. C. In this composite from a B6 heterozygote, the TM has not completely developed and trabecular beams are absent. Cells with large nuclei (inset, arrowheads) occupy the normal location of the TM and resemble the mesenchyme from which the TM is normally derived. SC is present and of normal length, but compressed (arrowheads). The iris is attached to the peripheral cornea, creating a false angle, covered with cells resembling corneal endothelial cells and with acellular tissue resembling Descemet’s membrane. D. In this heterozygote, SC appears normal, but the TM is slightly hypoplastic. E. SC and TM are absent and the iris is attached to the cornea. Schwalbe’s line is enlarged (arrow). F. SC and TM (arrows) are approximately half length (see B). G. Normal SC and TM are absent and there is an extensive iridocorneal adhesion. Original magnification × 400, except for inset, × 630. (From Smith, R., Hum. Mol. Genet., 9:1021–1032, 2000. With permission of Oxford University Press.)

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FIGURE 8.26 Iridocorneal angle defects. The strain abbreviations are the same as in Figure 8.24, with the addition of 129BS, a strain of mixed 129 and Black Swiss background. A. SC is absent and a large blood vessel (V) is attached to a hypoplastic TM. B. In this region, SC and TM are relatively normal, reflecting the variation noted in different ocular locations. C. SC and TM are absent and there is an enlarged Schwalbe’s line (arrow). D. SC and TM are absent and a short, deformed, clublike iris (resembling aniridia) is attached to the cornea. The ciliary body is hypoplastic (arrow). E. SC is short (arrowheads) and the trabecular beams are hypoplastic. A ciliary process is located abnormally at the posterior termination of the pars plana (arrow). F. SC and TM cannot be identified. G. The TM and SC are relatively normal in this region of a double heterozygote. H. SC and TM are absent and there is a long iridocorneal attachment. The corneal inflammatory infiltrate is related to the presence of open eyelids at birth. The ciliary body is malformed (arrow). Original magnification × 400. (From Smith, R., Hum. Mol. Genet., 9:1021–1032, 2000. With permission of Oxford University Press.)

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Although found in all backgrounds (129 Black Swiss, 129/SvEvTac, C57BL/6J, and CHMU/Le) using histological techniques, the highest frequency of visible abnormalities are demonstrated in Foxc1+/– mice on a C57BL/6J background, in CHMU/Le (Foxc1ch/+) mice, and in double heterozygotes (Foxc1+/–Foxc2+/–). Slit lamp evaluation in mice of the same strain and genetic background demonstrates variations between eyes of individuals. The variation in clinical phenotypes is reflected in the morphological phenotypes. An individual eye typically contains both normal and abnormal regions. This observation and the knowledge that intraocular pressure can remain normal with a limited amount of normal trabeculum147,148 may explain the normal intraocular pressure observed in these mice. The appearance of the trabecular meshwork and Schlemm’s canal is closely linked. In affected eyes, when the trabecular meshwork is hypoplastic or absent there is usually a small or absent Schlemm’s canal. When the trabecular meshwork is normal, Schlemm’s canal is also normal. Iris strands that attach to the cornea are frequently identified. In some severely affected mice, there are peripheral anterior synechias and occasional peripheral endothelialization of the iris surface. Especially in Foxc1ch/+ mice, there is enlargement of the termination of Descemet’s membrane (Schwalbe’s line) and the iris is frequently short and deformed, resembling aniridia. The ciliary body is hypoplastic and some processes are aberrantly located at the retinal end of the pars plana in Foxc1ch/+ heterozygotes (Figures 8.25 through 8.27). Electron microscopy confirms the findings of light microscopy and demonstrates that collagen fibers at the corneoscleral junction are half the diameter of those in wild-type mice.135

MICROPHTHALMIA AND ANOPHTHALMIA Absence (anophthalmia) or reduction in size (microphthalmia) of the eye are uncommon congenital abnormalities found in many animals, including humans,72 dogs,161 cats,162 pigs,163 hamsters,164 guinea pigs,165 and rats.166,167 Microphthalmia is a specific phenotypic feature of several spontaneous or radiation-induced mouse mutations including fidget (Fignfi),168,169 the many variants at the microphthalmia

FIGURE 8.27 Schlemm’s canal and trabecular meshwork abnormalities in Foxc1 mutant mice. All images are of strain 129 mice of the indicated genotypes. Except for F which is close to the iris root, the remaining figures are taken from the middle portion of SC. A. In a wild-type eye, SC is lined with endothelium and contains giant vacuoles (V). Several well-formed trabecular beams are present. B. SC is present and lined by thin endothelium. Abnormal tissue lacking the structure of normal trabecular beams separates the anterior chamber (AC) from SC. C. In this wild-type mouse, SC has a robust endothelial lining (E). The juxtacanalicular TM has a normal beam structure consisting of trabecular cells (arrows) surrounding densely packed collagen and elastic tissue. D. In this affected juxtacanalicular region, SC has a thin endothelial lining and there is little collagen or elastic tissue, compared with C. This region is from the same eye shown in B, in a nearby location. E. The TM adjacent to the anterior chamber has a normal appearance. However, adjacent to SC, the TM is dense and lacks intertrabecular spaces. F. In this composite view, the endothelial lining of SC is continuous, but lacks giant vacuoles. Normally, five to six giant vacuoles would be expected in a similar area of a normal mouse. G. Normal TM is completely absent in this heterozygote. A cell type not normally found in this area lines the anterior chamber (AC). These cells may represent mesenchyme that has not undergone normal development. H and I. Higher magnification of the corneoscleral transition zone above SC from a wild-type eye (in H) and a heterozygote (in I). The collagen bundles of the wild-type mouse are about twice the diameter of the heterozygote. J. The iris (I) and AC close to the iris root are normal. The corneoscleral transition zone that typically has little pigment is abnormal. The abnormal tissue separating the corneoscleral junction from the AC includes collagen and dendritic melanocytes. SC and TM are absent, although they are normally present at this location. Original magnification approximately × 12,000, except H and I, × 30,000. (From Smith, R., Hum. Mol. Genet., 9:1021–1032, 2000. With permission of Oxford University Press.)

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locus (Mitf) (see Chapters 9 and 10),170-172 Pax6 mutations associated with small eye,173-176 and vacuolated lens with microphthalmia (Vlm).177 Several genetically engineered mice with microphthalmia have also been described.178-187 The references given represent only a partial listing. Finally, microphthalmia is found in ZRDCT/Ch mice188,189 and as a background lesion in inbred black mice, a finding described many years ago.190 Many different strains of black mice exhibit microphthalmia and the incidence may be as high as 10%.49,191,192 Local environmental factors193 and exogenous stimuli such as alcohol during pregnancy cause a marked increase in microphthalmia frequency. In fact, alcohol given around day 7 of pregnancy in C57BL/6J mice194-196 is useful as a mouse model of the fetal alcohol syndrome described in human offspring of mothers suffering from alcoholism.197,198 For unexplained reasons, females are affected 6.2 times as often as males and the right eye 5.8 times as often as the left.193,199 In a single mouse there is often marked asymmetry, with one normal and one affected eye or one microphthalmic and one anophthalmic eye. Careful orbital dissection in an “anophthalmic” mouse often demonstrates a very small eye or remnants of an eye. Not infrequently, an eye apparently normal on clinical examination will be slightly smaller than normal and contain typical intraocular malformations associated with microphthalmia on microscopic examination. Thus, the true incidence of both microphthalmia and anophthalmia depends on whether it is based on casual clinical examination, slit lamp evaluation, or microscopic examination.

FIGURE 8.28 A. Anophthalmia in C57BL/6J mice. The eyelids (L) surround an empty conjunctival sac (C) that contains no eye. Orbital structures including the Harderian gland (H) are present. Original magnification × 50. B. In a microphthalmic C57BL/6J eye, the lens is cataractous and there has been intraocular extrusion of lens material (*). A dysplastic rosette (arrow) is associated with incomplete retinal differentiation. There is detachment of the retina. The vitreous contains undifferentiated pigmented cells. Original magnification × 100. C and D. Coloboma of the optic nerve in Bst/+ mice. In C, there is gross malformation of the peripapillary retina (arrow). A portion of the retina (R) has extended into the cavity of the coloboma. No nerve fiber layer is identifiable and the optic nerve (ON) consists of undifferentiated fibrous and pigmented tissue. In D, both retina and choroidal tissue extend into the coloboma. Original magnification × 200. (A and B, from Smith, R., Lab. Animal Sci., 44:551–560, 1994. With permission.)

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In addition to the corneal opacities discussed earlier (see Figure 8.5) inbred black mice have other intraocular abnormalities. Cataracts are frequently present and often associated with keratolenticular adhesions. The cataracts are usually cortical, but lens extrusion may develop in the posterior lens capsule. Multiple foci of dysplastic retina are common in microphthalmic eyes. In more severely affected eyes, the anterior chamber is absent and the iris and ciliary body severely malformed. In these eyes the cornea is often thickened and hypercellular, most likely due to loss or failure of development of the corneal endothelium. In true anophthalmia, no ocular remnants can be identified and the empty orbit is lined by thin conjunctiva. Other orbital structures, such as Harderian and lacrimal glands, are usually present in their normal locations. Severely microphthalmic and anophthalmic eyes are also at risk for recurrent infections because epithelial and environmental debris stagnate in the empty conjunctival sac, providing a nidus for infection by agents such as Corynebacteria spp. (Figure 8.28).49 Ocular changes in microphthalmia may vary in degree and extent of involvement of specific ocular structures, but for the most part are similar to those described in inbred black mice. An example of a specific difference are colobomas of the optic nerve and retina (see Chapter 11) commonly seen in mutations at the microphthalmia locus (Mitf),200 in Bst+/– mice,201 in Pax21Neu mice,202 in mice with the coloboma deletion (Cm),203 and in the extra toes (Gli3Xt) mutation (see Figure 8.20).204,205 These examples are a reminder that similar clinical and morphological phenotypes may be produced by completely different genes.

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149. Bejjani, B.A. et al., Mutations in CYP1B1, the gene for cytochrome P4501B1, are the predominant cause of primary congenital glaucoma in Saudi Arabia, Am. J. Hum. Genet., 62:325, 1998. 150. Bejjani, B.A. et al., Multiple CYP1B1 mutations and incomplete penetrance in an inbred population segregating primary congenital glaucoma suggest frequent de novo events and a dominant modifier locus, Hum. Mol. Genet., 9:367, 2000. 151. Andersen, J.S. et al., A gene responsible for the pigment dispersion syndrome maps to chromosome 7q35-q36, Arch. Ophthalmol., 115:384, 1997. 152. Shields, M.B., Progressive essential iris atrophy, Chandler’s syndrome and the iris nevus (Cogan Reese) syndrome: a spectrum of disease, Surv. Ophthalmol., 24:3, 1979. 153. Silver, L.M., Mouse Genetics: Concepts and Applications, Oxford University Press, New York, 1995. 154. Waring, G.O., Rodrigues, M.M., and Laibson, P.R., Anterior chamber cleavage syndrome: a stepladder classification, Surv. Ophthalmol., 20:3, 1975. 155. Shields, M.B., Axenfeld-Rieger syndrome: a theory of mechanism and distinctions from the iridocorneal endothelial syndrome, Trans. Am. Ophthamol. Soc., 81:736, 1983. 156. Shields, M.B. et al., Axenfeld-Rieger syndrome. A spectrum of developmental disorders, Surv. Ophthalmol., 29:387, 1985. 157. Nishimura, D.Y. et al., The forkhead transcription factor FKHL7 is responsible for glaucoma phenotypes which map to 6p25, Nat. Genet., 19:140, 1998. 158. Mears, A.J. et al., Mutations of the forkhead/winged helix gene, FKHL7, in patients with AxenfeldRieger anomaly, Am. J. Hum. Genet., 63:1316, 1998. 159. Kume, T. et al., The Forkhead/Winged helix gene Mf1 is disrupted in the pleiotropic mouse mutation congenital hydrocephalus, Cell, 93:985, 1998. 160. Hong, H., Lass, J.H., and Chakravarti, A., Pleiotropic skeletal and ocular phenotypes of the mouse mutation congenital hydrocephalus (ch/Mf1) arise from a winged helix/forkhead transcription factor gene, Hum. Mol. Genet., 8:625, 1999. 161. Zhang, R.L. et al., Analysis of eye lens-specific genes in congenital hereditary cataracts and microphthalmia of the miniature schnauzer dog, Invest. Ophthalmol. Vis. Sci., 32:2662, 1991. 162. Robson, J.A., Organization of the dorsolateral geniculate nucleus in a cat with congenital microphthalmia, Brain Res., 430:81, 1987. 163. Szabo, K., Congenital Malformations in Laboratory and Farm Animals, Academic Press, San Diego, 1989. 164. Wilson, J.G. and Fraser, F.C., Eds., Handbook of Teratology, Plenum Press, New York, 1977. 165. Komich, R.J., Anophthalmos: an inherited trait in a new stock of guinea pigs, Am. J. Vet. Res., 32:2099, 1971. 166. Rao, S.H. and Sesikeran, B., Congenital anophthalmia in CFY rats: a newly identified autosomal recessive mutation, Lab. Animal Sci., 42:623, 1992. 167. Wyse, J.P.H. and Hollenberg, M.J., Complicated colobomatous microphthalmos in the BW rat: a new form of inherited retinal degeneration, Am. J. Anat., 149:377, 1977. 168. Gruneberg, H., Two new mutant genes in the house mouse, J. Genet., 45:22, 1943. 169. Truslove, G.M., The anatomy of the development of the fidget mouse, J. Genet., 54:64, 1956. 170. Hodgkinson, C.A. et al., Mutations at the mouse microphthalmia locus are associated with defects in a gene encoding a novel basic-helix-loop-helix-zipper protein, Cell, 74:395, 1993. 171. Steingrimsson, E. et al., Molecular basis of mouse microphthalmia (mi) mutations helps explain their developmental and phenotypic consequences, Nat. Genet., 8:256, 1994. 172. Yajima, I. et al., An L1 element intronic insertion in the black-eyes white (Mitfmi-bw) gene: the loss of a single Mitf isoform responsible for the pigmentary defect and inner ear deafness, Hum. Mol. Genet., 8:1431, 1999. 173. Hogan, B. et al., Small eye (Sey): a mouse model for the genetic analysis of craniofacial abnormalities, Development, 103:115, 1988. 174. Barinaga, M., Focusing on the eyeless gene, Science, 267:1766, 1995. 175. Schedl, A. et al., Influence of PAX6 gene dosage on development: overexpression causes severe eye abnormalities, Cell, 86:71, 1996. 176. Collinson, J.M., Hill, R.E., and West, J.D., Different roles for Pax6 in the optic vesicle and facial epithelium mediate early morphogenesis of the murine eye, Development, 127:945, 2000.

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177. Kratochvilova, J., Dominant cataract mutations detected in offspring of gamma-radiated male mice, J. Hered., 72:302, 1981. 178. Breitman, M.L. et al., Genetic ablation: targeted expression of a toxin gene causes microphthalmia in transgenic mice, Science, 23:1563, 1987. 179. Landel, C.P. et al., Lens specific expression of recombinant ricin induces developmental defects in the eyes of transgenic mice, Genes Dev., 2:1168, 1988. 180. Breitman, M.L. et al., Analysis of lens cell fate and eye morphogenesis in transgenic mice ablated for cells of the lens lineage, Development, 106:457, 1989. 181. Klein, K.L. et al., Embryology and morphology of microphthalmia in trangenic mice expressing a gamma F-crystallin/diphtheria toxin A hybrid gene, Lab. Invest., 67:31, 1992. 182. Griep, A.E. et al., Tumorigenicity by human papillomavirus type 16 E6 and E7 in transgenic mice correlates with alterations in epithelial cell growth and differentiation, J. Virol., 67:1373, 1993. 183. Kastner, P. et al., Genetic analysis of RXRα developmental function: convergence of RXR and RAR signaling pathways in heart and eye morphogenesis, Cell, 78:987, 1994. 184. Egwuagu, C.E. et al., Ectopic expression of gamma interferon in the eyes of transgenic mice induces ocular pathology and MHC Class II gene expression, Invest. Ophthalmol. Vis. Sci., 35:332, 1994. 185. Matsuo, I. et al., Mouse Otx2 functions in the formation and patterning of rostral head, Genes Dev., 9:2646, 1995. 186. Dattani, M.T. et al., Mutations in the homeobox gene HESX1/Hesx1 associated with septo-optic dysplasia in human and mouse, Nat. Genet., 19:125, 1998. 187. Robinson, M.L. et al., Disregulation of ocular morphogenesis by lens-specific expression of FGF-3/Int2 in transgenic mice, Dev. Biol., 198:13, 1998. 188. Silver, J. and Hughes, A.F.W., The relationship between morphogenetic cell death and the development of congenital anophthalmia, J. Comp. Neurol., 157:281, 1974. 189. Harch, C., Chase, H.B., and Gonsalves, N.I., Studies on an anophthalmic strain of mice. VI. Lens and cup interaction, Dev. Biol., 63:352, 1978. 190. Chase, H.B., Studies on an anophthalmic strain of mice. III. Results of crosses with other strains, Genetics, 27:330, 1942. 191. Kalter, H., Sporadic congenital malformations of newborn inbred mice, Teratology, 1:193, 1968. 192. Smith, R.S. and Sundberg, J.P., Ophthalmic abnormalities in inbred mice, in Pathobiology of the Aging Mouse, Mohr, U. et al., Eds., ILSI Press, Washington, D.C., 1996, 117. 193. Pierro, L.J. and Spiggle, J., Congenital eye defects in the mouse. II. The influence of litter size, litter spacing, and suckling of offspring on risk of eye defects in C57BL mice, Teratology, 2:337, 1969. 194. Sulik, K.K., Johnston, M.C., and Webb, M.A., Fetal alcohol syndrome: embryogenesis in a mouse model, Science, 214:936, 1981. 195. Webster, W.S. et al., Some teratogenic properties of ethanol and acetaldehyde in C57BL/6J mice: implications for the study of the fetal alcohol syndrome, Teratology, 27:331, 1983. 196. Cook, C.S., Nowotney, A.Z., and Sulik, K.K., Fetal alcohol syndrome: eye malformations in a mouse model, Arch. Ophthalmol., 105:1576, 1987. 197. Chan, T. et al., Ocular manifestations in fetal alcohol syndrome, Br. J. Ophthalmol., 75:524, 1991. 198. Hellstrom, A., Svensson, E., and Stromland, K., Eye size in healthy Swedish children and in children with fetal alcohol syndrome, Acta Ophthalmol., 75:423, 1997. 199. Tyan, M.L., Effects of H-2 and vitamin A on eye defects in congenic mice, Proc. Soc. Exp. Biol. Med., 199:123, 1992. 200. Scholtz, C.L. and Chan, K.K., Complicated colobomatous microphthalmia in the micropthalmic (mi/mi) mouse, Development, 99:501, 1987. 201. Smith, R.S. et al., The Bst locus on mouse chromosome 16 is associated with age-related subretinal neovascularization, Proc. Natl. Acad. Sci. U.S.A., 97:2191, 2000. 202. Favor, J. et al., The mouse Pax21Neu mutation is identical to a human PAX2 mutation in a family with renal-coloboma syndrome and result in developmental defects of the brain, ear, eye, and kidney, Proc. Natl. Acad. Sci. U.S.A., 93:13870, 1996. 203. Hes, E.J. et al., Deletion map of the Coloboma (Cm) locus on mouse chromosome 2, Genomics, 21:257, 1994.

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204. Franz, T. and Besecke, A., The development of the eye in homozygotes of the mouse mutant extra-toes, Anat. Embryol., 184:355, 1991. 205. Pohl, T.M., Mattei, M., and Ruther, U., Evidence for allelism of the recessive insertional mutation add and the dominant mouse mutation extra-toes (Xt), Development, 110:1153, 1990.

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Choroid, Lens, and Vitreous Richard S. Smith

CONTENTS Choroid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .162 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .162 Development Defects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .162 Retinol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .162 Platelet Storage Pool Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .164 Microphthalmia in Mitf Locus Mutations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .165 Uveitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .168 Trauma and Its Sequelae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .169 Vascular Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .169 Induced Chorioretinal Neovascularization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .171 Spontaneous Choroidal Neovascularization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .171 Neoplasms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .171 Lymphoproliferative Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .171 Retinoblastoma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .173 Pigmented Neoplasms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .173 Choroidal Degeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .175 Lens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .175 Cataract Types . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .176 Cataract Morphology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .177 General . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .177 Capsular-Epithelial . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .178 Nuclear and Cortical . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .178 Cortical Liquefaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .178 Lens Extrusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .180 Neoplasia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .182 Cataracts in Ocular and Systemic Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .182 Vitreous . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .182 Developmental Abnormalities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .182 Vitreous Agenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .182 Persistent Hyperplastic Primary Vitreous . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .184 PHPV in p53-Null Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .185 PHPV in Norrie’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .185 PHPV with Ectoptic Gene Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . .185 PHPV Induced by Retinoic Acid and Its Receptors . . . . . . . . . . . . . . . . . . .185 Vitreous Hemorrhage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .187 Vitreous Inflammation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .187 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .188

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CHOROID INTRODUCTION Unlike the anterior uvea (iris and ciliary body) which has a dual origin from neuroepithelium and mesenchyme, the choroid is derived solely from mesenchyme including neural crest. Its principal physiological function is to serve the metabolic needs of the outer retina. The retina, retinal pigment epithelium, and choroid function almost as a single unit because of their close anatomic and functional interactions. As such, it is often necessary to discuss both structures when reviewing the effects of a specific disease. To keep the major discussion of a disease process in one location, the editors have divided retinal and choroidal diseases, based on what they consider the structure most importantly affected. Dendritic melanocytes and vascular channels occupy most of the choroid. After melanocytes and endothelial cells, mast cells are the most abundant choroidal cell, although their role in this location remains undefined.1 Macrophages, lymphocytes, and ganglion cells of the autonomic nervous system are frequently identified in normal choroidal tissue. Bruch’s membrane lies between choroid and retina and contains components derived from both the choroid and the retinal pigment epithelium.

DEVELOPMENTAL DEFECTS Retinol The effects of maternal vitamin A (retinol) deficiency on fetal development have been known for nearly 50 years. Failure of lid closure, cataracts, colobomas, persistent hyperplastic primary vitreous (see below), and abnormal folding of the sensory retina were originally reported in vitamin A–deficient rats.2 Retinoic acid is the active derivative of retinol and exerts the broadest effects during development. The complex family of retinoid receptors (RAR) includes RARα, β, γ, and their isoforms as well as RXRα, β, and γ (reviewed in Lohnes3). Gene-targeted mice lacking various combinations of RARs and RXRs have been generated and reveal a broad range of developmental defects. All groups of mice die at birth or shortly thereafter, with the exception of single mutants with targeted mutations in either Rarα1 or Rarβ2. In double mutants, there are malformations of the eye, heart, aortic arches, kidney, lungs, trachea, and skeleton.3-6 In addition to potential orbital bony defects, agenesis of the Harderian gland, open eyelids, and agenesis of the nasolacrimal drainage system, RAR double mutants demonstrate numerous intraocular defects including microphthalmia. The most widely affected mutant genotypes are the αγ and β2γ double mutants. Cornea, conjunctiva, anterior chamber, and lens may all be absent. Presence of a persistent keratolenticular stalk is likely a reflection of the C57BL/6 background utilized for creation of the null mutants.7 Chorioretinal and optic nerve colobomas occur in all mice of these two genotypes. In regions where the choroid was absent, the overlying sclera was also thinner than normal. A central feature in αγ and β2γ double mutants is the presence of persistent hyperplastic primary vitreous (see below).3,6 Similar findings were also demonstrated in the small number of RXRα null mutants that survive to E16.5 and in different combinations of haploinsufficient genotypes of RAR and RXR isoforms (Figure 9.1).5 Ocular abnormalities also occur in mice exposed to excessive amounts of retinoic acid during development. When pregnant mice are given intraperitoneal retinoic acid at gestation day 7, nearly all offspring develop microphthalmia. In about one third, failure of closure of the fetal fissure and persistent hyperplastic primary vitreous were observed. The lens was absent in a few mice and ruptured in others. Corneolenticular adhesions were common and about 20% of the mice had abnormal formation of the chamber angle structures.8 These findings have been reported as background lesions in C57BL/6 mice,7 although at a much lower incidence (see Chapter 4). An increased frequency of

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FIGURE 9.1 RAR null mutants. Comparison of frontal sections of the eye at E18.5. A and E, wild-type; B to D, F1 RAR double mutant mice. The genotype is indicated in each micrograph. Abbreviations: A, anterior chamber; C, corneal stroma; EP, corneal epithelium; ER, everted retina; F, fibrous retrolenticular membrane; I, iris; IR, internal leaf of retina; J, conjunctival sac; L, lens; ON, optic nerve; OR, outer leaf of the retina; PO, periocular mesenchyme; C, vitreous; Y, lids; asterisks, cartilaginous nodules replacing ethmoid bone. In G, the large arrow points to the persistent corneolenticular stalk; in I, the arrow points to a cartilaginous nodule in the retrolenticular membrane. In all mutant eyes there is a coloboma of the optic nerve and retina, the cornea is hypoplastic and the anterior chamber is generally absent. (From Chambon, P. et al., Development, 120:2723–2748, 1994. With permission of the Company of Biologists, Ltd.)

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microphthalmia-associated lesions has also been reported after fetal exposure to alcohol at the same gestational time period.9 Platelet Storage Pool Disease Organelle defects in lysosomes, melanosomes, and platelet granules underlie the albinism, coagulation defects, and abnormal lysosomal function characteristic of the human diseases Chediak–Higashi (CHS) and Hermansky–Pudlak syndromes (HPS).10-13 The mouse orthologue of the gene mutated in CHS, the lysosomal trafficking regulator (Lyst) has mutant recessive alleles such as the original beige allele (Lystbg), which cause lysosomal trafficking defects in addition to a lighter coat color. There are 14 mouse models of HPS, reflecting the genetic heterogeneity of this syndrome (reviewed in Gwynn et al.14). All mice with storage pool disease are characterized by pale coat colors. The decrease in uveal pigmentation produces similar clinical phenotypes, although the appearance of eyes in Lystbg/Lystbg mice is distinct from that of HPS model strains. Beige mice exhibit iris transillumination as soon as their eyes open, but large globular regions of dark pigment interrupt the reflected light. In contrast, the HPS strains have diffuse, marked iris transillumination at an early age that is nearly as prominent as that seen in a true albino mouse (Figure 9.2). The morphological phenotype of Lystbg/Lystbg mice correlates with their clinical appearance. The iris, ciliary body, choroid, and retinal pigment epithelium of these mice and related strains, such as SB/Le, contain giant melanosomes that often clump together (Figure 9.1).15-17 These large aggregations explain the patchy transillumination observed in these mice. Similar collections of clumped melanosomes also occur in pallid (Pldnpa/Pldnpa) mice.18 In addition, there may be an overall decrease in total melanosome numbers.16 In the retinal pigment epithelium, the presence of giant secondary phagolysosomes suggests a failure of the normal process of outer segment digestion.19 Free melanocytes, possibly macrophages laden with pigment and clumps of free melanosomes, aggregate in the anterior chamber. These are likely derived from iris melanocytes or iris pigment epithelial cells that rupture, contributing to the iris atrophy that is a constant feature of this model. The choroid is thickened by accumulations of melanocytes congested with giant melanosomes. In SB/Le mice, mineralized iris cysts are a common feature usually found on the posterior iris surface (Figures 9.2 and 9.3). Although difficult to photograph, with sufficient iris atrophy they can be visualized with the high power of the slit lamp. The 14 currently reported mouse models of HPS involve multiple mutations on eight different chromosomes while mutations responsible for human HPS homologues are located on ten different chromosomes. This observation reflects the complexity of molecular pathways that terminate in similar clinical phenotypes.20 The ocular findings in HPS are principally pigmentary abnormalities. Two HPS models, gunmetal (gm) and reduced pigment (rp) were reported to be lacking ocular pigmentary disturbances.20-22 However, these reports were limited to clinical observations. Light and transmission electron microscopy of rp/rp mice demonstrates decreased pigmentation of both the choroid and retinal pigment epithelium (Smith and Peters, unpublished observations) (see Figures 9.2 and 9.3). Although there is variation in ocular features in different HPS models, all pigmentary changes result from a decrease in either total melanosome number or in amount of melanin deposited within the melanosome. Melanosomes tend to be smaller than normal in contrast to the large pigment clumps typical of CHS model mice. Melanosome number is severely decreased and most remaining melanosomes are arrested at either stage I or II of pigment deposition.23,24 Abnormal melanosomes are evident in all parts of the uvea including iris, ciliary body, and choroid (Smith and Peters, unpublished observation) as well as the retinal pigment epithelium (Figure 9.3) (reviewed in Swank et al.20).25 In addition, the normally prominent basal infoldings of the retinal pigment epithelial cells are markedly reduced, a finding that is most conspicuous in the pearl strain (Ap3b1pe/Ap3b1pe).16 The remaining ocular structures are normal.

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FIGURE 9.2 Platelet storage pool deficiency in SB/Le mice. A. At age 6 months, iris pigment is both sparse and coarse, allowing prominent transillumination of the iris. B. Iris and ciliary body of 13-day-old mouse. Pigment granules in both ciliary (arrow) and iris epithelium are larger than normal. Original magnification × 400. C. By 4 months of age, large, pigment-filled cells are found in the anterior chamber (arrow) and in the root of the iris (arrowhead) and trabecular meshwork. Original magnification × 400. D. In older mice, unusually large pigment granules are found in both iris stroma (arrow) and pigment epithelium (arrowhead). Original magnification × 200. E. In older mice, particularly females, the iris becomes atrophic and cystic structures filled with basophilic material (*) develop on the posterior surface of the iris. Original magnification × 200. F. Although the material in the cystic structures shatters and is lost during sectioning, a von Kossa stain demonstrates that the basophilic material represents focal mineralization (arrow). Original magnification × 400.

Microphthalmia in Mitf Locus Mutations Albinism or coat color dilution also occurs in mice with mutations of the microphthalmia-associated transcription factor (Mitf). Numerous allelic mutations of this gene have been reported and many have microphthalma, ocular and coat color abnormalities, and osteopetrosis. Clinical phenotypes vary for the different alleles and there are complex interactions between the alleles.25 Although the

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FIGURE 9.3 Platelet storage pool deficiency. A. A fundus photograph of a 9-month-old SB/Le mouse demonstrates irregular pigmentation of the choroid and RPE. The optic nerve is pale and vessels are difficult to see, because this strain is homozygous for Pde6brd1. B. The choroid and retinal pigment epithelium of a B6.Cg-Lystbg/Lystbg mouse contains melanin granules that are much larger than normal. Original magnification × 630. C. A cross between C57BL-rp/+ and R26 has pigment granules of normal size, but there is focal depigmentation of the RPE (arrow). Original magnification × 630. D. In cappucino mice (cno/cno) there is nearly complete loss of melanin in both the choroid (C) and retinal pigment epithelium (arrow). The retina (R) of this strain is abnormal, as it is homozygous for the Pde6brd1 allele. E. In a normal cno/+ mouse, the retinal pigment epithelium (PE) and the choroid (C) contain numerous melanin granules. Original magnification × 5000. F. In a cno/cno mouse, there are only a few very small, poorly melanized melanosomes (arrows) in the RPE. Choroidal melanin granules (inset, arrow) are also extremely small and incompletely melanized. Original magnification × 5000; inset × 24,000. (C to F courtesy of Dr. Luanne Peters.)

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eyes of mice with a variety of Mitf alleles are smaller than normal, they lack many lesions characteristic of microphthalmic C57BL/6 mice (see Chapter 4). Colobomas of the optic nerve, choroid, and retina (see Chapter 11) are typical of mutations at the microphthalmia locus.26-29 Although normal choroidal pigmentation may develop, shortly after birth melanosomes fragment and release pigment that is ingested by phagocytic cells (pigment dispersion and incontinence).30 With other Mitf alleles, melanosomes may be absent from the choroid (reviewed in Hodgkinson et al.28). In mice with a random DNA insertional mutation at the Mitf locus, choroidal melanocytes were totally absent, suggesting a failure of neural crest migration (see Figure 9.4).31 A similar clinical phenotype is seen in patients with Waardenburg syndrome type 2, probably due to a mutation at the MITF locus.32

FIGURE 9.4 Microphthalmia alleles. A. The eye of a 22-day-old B6C3Fe-a/a-Mitfmi/Mitfmi mouse demonstrates multiple malformations and is only about half normal size. There is a diffuse cataract (C) and only minimal retinal differentiation. Original magnification × 50. B. In the anterior segment the ciliary body is absent and a malformed iris (I) adheres to the cataractous lens (L) and to the hypercellular cornea (C). The peripheral retina (R) is attached to the lens capsule. Original magnification × 200. C. In a C57BL/6J-Mitfmi-vit/Mitfmi-vit mouse, there is pigment dispersion with free macrophages in the anterior chamber (arrow). Original magnification × 100. D. The iris structure in a mouse homozygous for Mitfmi-vit is completely normal. In the same eye, the choroid at the back of the eye contains no melanin (arrow) and the RPE (arrowhead) has decreased pigment. F. In the anterior portion of the choroid of the same mouse as in E, choroidal pigmentation is normal, although the RPE is hypopigmented. D to F original magnification × 630.

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UVEITIS Inflammation of iris, ciliary body, or choroid is referred to as uveitis. In human eyes, uveitis is associated with a broad range of etiologies that include infectious disease (viral, bacterial, and fungal), immunodeficiency, autoimmunity, immune complex deposition, post-traumatic, and with associated systemic diseases such as lupus and periarteritis.33 In many cases, despite intensive medical evaluation, the etiological agent remains unknown.33,34 Whereas uveitis in humans frequently occurs spontaneously due to one of these causes, uveitis in mice is usually induced by the investigator, nearly always through immune mechanisms. In most cases of uveitis, the inflammation is not limited to the pigmented layers of the eye and often extends to involve the retina, vitreous, sclera, cornea, and lens. Although the cytokine interferon-γ (IFN-γ) is not normally found in the eye, it is expressed as a response to different inflammatory stimuli (reviewed in Geiger et al.35). Mice that express IFN-γ in the retina develop acute intraocular inflammation associated with peripheral keratitis, cataract formation, anterior chamber pigment dispersion, retinal perivasculitis, and severe photoreceptor destruction on a background of diffuse uveitis.35 The degree of this response is not surprising, since many macrophages and dendritic cells are found in the mouse uveal tract.36 Diffuse ocular inflammation is characteristic of experimental autoimmune uveoretinitis (EAU). EAU is initiated by immunization with either retinal S-antigen or retinol-binding protein 3, interstitial (RBP3, formerly, interphotoreceptor retinoid-binding protein, or IRBP) and mediated by T lymphocytes. The disease appears about 3 weeks after immunization. Inflammation is severe with scleritis, uveitis, retinitis, focal retinal detachment, focal photoreceptor dropout, retinal perivasculitis, and subretinal neovascularization (Figure 9.5).37,38 There are many similarities between EAU and

FIGURE 9.5 Experimental autoimmune uveoretinitis. A. Normal control mouse retina. B. In an early mild case, numerous lymphocytes are present in the vitreous (arrowhead). A subretinal granuloma has extended into both inner and outer nuclear layers (arrow). C. In a more severe case, the retina is edematous and folded (F) and there is an intense inflammatory infiltrate most prominent in the choroid (arrow), but also involving the RPE. Inflammatory cells are also present in the retina, particularly in the inner plexiform layer. D. A severe case manifests all the features of the mild and moderate inflammatory response. There is more frequent subretinal granuloma formation and large areas of photoreceptors have been destroyed. The arrow indicates remaining photoreceptor nuclei. Original magnification × 200. (Photographs courtesy of Dr. Rachel Caspi.)

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human diseases such as sympathetic ophthalmia, Vogt-Koyanagi-Harada syndrome, and Behcet’s disease.37,39 Both antigen-presenting dendritic cells and nonantigen-presenting macrophages from the choroid36 have been implicated in EAU.39 Systemic and intravitreal endotoxin administration (bacterial lipopolysaccharide) produces an acute uveitis that is most severe in the iris and ciliary body and less so in the choroid. Within a few hours of injection, neutrophils appear in iris, ciliary body, aqueous, and vitreous. By 9 h, lymphocytes and mononuclear phagocytic cells are abundant.34,40,41 Inflammation peaks by 24 h and resolves within 48 h without permanent consequences. Genetic factors likely play an important role in the response to this stimulus. Even closely related strains vary in their response, emphasizing the critical importance of genetic background in interpreting experimental results (see Chapters 4 and 5). For example, C3H/HeN mice demonstrate a marked response, whereas C3H/HeJ mice fail to develop uveitis.41,42 Mice have been used as experimental models for induction of ocular herpes simplex disease. When live virus is injected into the anterior chamber, two different types of uveitis result. In the injected eye, there is an intense acute inflammatory reaction that involves cornea, iris, and ciliary body with sparing of the posterior ocular structures. In the uninjected eye, after 1 to 3 weeks, a massively destructive process develops that includes vitritis, necrotizing retinitis, and intense choroiditis. The inflammatory infiltrate includes lymphocytes, plasma cells, and multinucleated giant cells.43 A similar outcome occurs after intercerebral or intranasal viral inoculation.44

TRAUMA AND ITS SEQUELAE Most ocular injuries in mice are superficial corneal wounds from cage mates or bedding materials. Although uncommon, it is possible to injure the eye in the process of blood collection from the orbital venous sinus. The sclera is thin and easily punctured if the collecting tube grazes the eye. Although not always evident at the time of injury, the eye is likely to become swollen and exophthalmic because of intra- and extraocular hemorrhage. Extensive choroidal and retinal damage are common sequelae. In human eyes, the occurrence in the noninjured eye of intense inflammation is known as sympathetic ophthalmia.33 A similar reaction may occur in mice, although the extent of inflammation is much less. After surgical placement of a small intravitreal ferrous foreign body, mild infiltration of T lymphocytes and macrophages develops in the unoperated eye during the first 2 weeks after surgery in mice.45 It is not clear if such a reaction might occur with lesser manipulation of one eye, but investigators should be aware of the possibility when using one eye as an untreated control. Ocular trauma or severe inflammation is likely to damage most intraocular structures. A common sequel to either event is atrophy, shrinkage, and disorganization of the globe, referred to as phthisis bulbi (Figure 9.6). Typical findings include thickening of the cornea and sclera, irregular shape of the eye, fibrosis of the vitreous and anterior chamber, mineralization of the lens and retina, and deposition of metaplastic cartilage or bone in the choroid. In the process, it becomes difficult, if not impossible, to identify intraocular structures such as iris, ciliary body, or retina.

VASCULAR DISEASE Sickle cell anemia in human patients results from a mutation in the hemoglobin beta gene that leads to production of a structurally abnormal hemoglobin. Erythrocytes assume a sickle shape and undergo accelerated destruction. The abnormal erythrocytes block capillaries, leading to vascular occlusion and focal ischemia. Both choroid and retina may be affected by post-ischemic vasoproliferation. Pigmented retinal spots commonly seen in sickle cell chorioretinopathy are caused by migration of retinal pigment epithelial cells into the retina (reviewed in Lutty et al.46). Of transgenic

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FIGURE 9.6 Phthisis bulbi. A. This phthisical eye is about one half normal diameter. The cornea (C) is thickened and irregular. The iris and ciliary body cannot be identified. The lens (L) is cataractous, displaced from its normal position and adherent to the retina (R). Original magnification × 50. B. The retina (R) is detached and missing most normal retinal layers. At the arrow, both choroid and retina have been replaced by a thick fibrovascular scar. Original magnification × 200. C. The thickened vascularized cornea is folded, particularly facing the anterior chamber where multiple folds in Descemet’s membrane (arrow) are evident. Original magnification × 100. D. In severely phthisical eyes, bone and cartilage may form, especially in the posterior portion of the eye. Here, retina and RPE have been replaced by well-differentiated cartilage (arrow). The choroid (C) is completely degenerated and the sclera (S) is thickened. Original magnification × 400.

mice that carry various constructs of the sickle cell allele of human hemoglobin β, 30% develop a morphological phenotype after age 15 months that resembles the human disease.46 Findings include focal retinal vascular occlusions associated with areas of retinal nonperfusion and the development of retinal arteriovenous anastamoses. Intra- and extraretinal neovascularization is observed in nonperfused regions. Perivascular migration of retinal pigment epithelial cells surrounds both retinal and choroidal vessels including areas of choroidal neovascularization.46 Older mice also develop severe photoreceptor loss that was attributed to occlusion of the choriocapillaris by fibrin and erythrocyte aggregation.47 Lupus erythematosis (SLE), a human disease with severe diffuse systemic effects, can also lead to ocular lesions including dry eyes, peripheral corneal melting, retinal vasculitis, and chorioretinopathy.33,48 Clinical and morphological phenotypes that resemble SLE have been reported in several inbred mouse strains including NZB, NZW, MRL/MpJ-Tnfrsf6lpr, BXSB/MpJ, and intercross (NZW X BXSB) F1 male offspring.25,48 These mice develop multiple systemic lesions including thrombocytopenia, lupus nephritis, and myocardial infarction. Small white retinal spots can be demonstrated on ophthalmoscopic examination. Choroidal arterioles are thickened and there is narrowing and occlusion of the choriocapillaris with thrombus formation.48 The relative ischemia this produces likely explains the loss of retinal pigment epithelium and photoreceptor destruction that follows.

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INDUCED CHORIORETINAL NEOVASCULARIZATION Degenerative changes in the macula (the retinal area responsible for the most acute vision) is the leading cause of blindness in developed countries in individuals over the age of 50.49,50 The neovascular form of macular degeneration has a particularly poor prognosis with 64% of patients losing most useful vision within 2 years of the onset of symptoms.51 The frequency of macular degeneration has led to development and characterization of several mouse models. Focal chorioretinal laser burns consistently stimulate the growth of new vessels in the choroid that extend through the laser-induced lesion and proliferate beneath the retina of mice.52,53 Similar morphological changes are produced by subretinal implantation of fibroblast growth factor 2–impregnated microspheres.54 Vascular endothelial growth factor (VEGF) is an angiogenic factor known to play an important role in ocular neovascularization.55,56 Chorioretinal neovascularization has also been produced in rats by adenoviral mediated expression of VEGF57 and in transgenic mice with the human VEGF gene regulated by the bovine rhodopsin promoter.58 In both models, new blood vessels develop in the choroid, proliferating through Bruch’s membrane beneath the retina or directly invading it. These morphological changes are similar to those that occur in spontaneous chorioretinal neovascularization (Figure 9.7).

SPONTANEOUS CHOROIDAL NEOVASCULARIZATION The molecular mechanisms responsible for human chorioretinal neovascularization remain obscure and are likely due to complex gene interactions.59,60 A choroidal neovascularization phenotype occurs in mice with the autosomal semidominant mutation belly spot and tail (Bst) on Chr 16 in the C57BLKS strain. This mutation is characterized by late closure of the choroidal fissure, coloboma of the optic nerve, and focal abnormalities of retinal differentiation.61 Choroidal neovascularization in Bst/+ mice develops between 7 and 12 months of age. The association between optic nerve coloboma and chorioretinal neovascularization is similarly observed in human eyes.62,63 In Bst/+ mice, 20% exhibit focal retinal detachments due to development of chorioretinal neovascularization that presents either as a focal plaque beneath the retina or extends into the retina. This abnormality is clearly visible on ophthalmoscopic examination. Fluorescein angiography reveals focal vascular leakage in areas with neovascularization. The retinal detachment is associated with the presence of pigmented phagocytic cells in the subretinal space and degeneration of photoreceptor outer segments. In addition, Bst/+ mice demonstrate colobomas of the optic nerve, focal retinal dysplasia, and patches of poorly differentiated retina61 (Figure 9.7).

NEOPLASMS Although mice remain one of the most important cancer models, relatively little has been written concerning ocular tumors. The cornea64 and anterior chamber65 have long been used to study tumor biology, tumor immunology,66 and angiogenesis67 because of the ability to observe tumor and vascular growth on a daily basis, without repeated surgical invasion. The experimental technology of such artificial systems is outside the scope of this book. Primary or metastatic ocular tumors are rarely seen in mice, but are likely overlooked due to lack of clinical or histopathologic evaluation.

LYMPHOPROLIFERATIVE DISORDERS The only systemic neoplasms that frequently involve the eye and orbit in mice are the lymphoproliferative disorders. These include not only polyclonal collections of lymphocytes, plasma cells, histiocytes, and polymorphonuclear leukocytes (which may be prelymphomatous lesions), but also monoclonal infiltrates of B cells, T cells, or histiocytes. Classification of lymphoproliferative lesions is complex and requires numerous specialized techniques that are thoroughly reviewed elsewhere.68,69

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FIGURE 9.7 Choroidal neovascularization. A. Fluorescein angiogram of a mouse with choroidal neovascularization taken 154 s after intraperitoneal injection. A large patch of fluorescein leakage is present temporal to the optic nerve (arrow). B. A vessel extends directly from the choroid (arrows) into the subretinal space, where it branches and extends into the outer nuclear layer of the retina (R). Original magnification × 400. C. A plaque of subretinal fibrovascular proliferation (P) has replaced the pigment epithelium and produced a focal retinal detachment (arrow). D. In this Bst/+ mouse, the choroidal neovascularization is associated with a coloboma of the optic nerve. Original magnification × 200. E. Immunodeficient mouse with disseminated lymphosarcoma. The anterior chamber contains many small malignant lymphocytes (arrow). Original magnification × 400. F. In the choroid (arrow), the same mouse demonstrates a dense infiltrate of neoplastic lymphocytes. Original magnification × 630.

Many spontaneous and induced mutations associated with lymphoma or leukemia are known, and the pattern of expression varies with different strain backgrounds. Lymphomas with involvement of many organs appear at higher frequency in some strains, such as BALB/c, SJL, DBA/2, C57BL, C58, NZB, and many transgenic strains.68,69 The choroid and orbit are the most frequent ocular sites invaded by lymphoma, although other eye tissues may be involved (illustrated in Chapter 11). The eye

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has not been reported as a primary site for lymphoma in mice, but may have been overlooked since primary ocular lymphoma occurs in human eyes.33

RETINOBLASTOMA In human eyes, retinoblastoma is the most common primary intraocular malignant neoplasm in infants and young children.33,70,71 Retinoblastoma can arise from mutations in the retinoblastoma gene (Rb) that acts in an autosomal dominant fashion with incomplete penetrance (reviewed in Windle et al.72 and Spencer33). Retinoblastoma in mice does not occur spontaneously, but has been produced in transgenic mice using the simian virus 40 T antigen driven either by the luteinizing hormone β-subunit gene72,73 or by the promoter of RBP3.74 Tumor morphology in these transgenic mice is similar to that seen in human retinoblastomas. The majority of the tumor cells are small and undifferentiated with hyperchromatic nuclei, scanty cytoplasm, poor cohesion, and a high mitotic index. Tumor necrosis and mineralization is common, often leaving perivascular cuffs of viable tumor cells. A subset of cells in mouse retinoblastomas undergoes photoreceptor differentiation and forms typical Flexner-Wintersteiner or Homer Wright rosettes.72 Although tumors arise in the retina, the lack of cohesion and the rapid growth rate leads to widespread invasion of other ocular structures including choroid, iris, ciliary body, cornea, vitreous, and optic nerve72,73 (Figure 9.8).

PIGMENTED NEOPLASMS Uveal melanoma is the most common primary malignant ocular tumor in human adults.33 Malignant pigmented intraocular tumors that metastasize and are lethal have been produced in mice.75,76 The majority of transgenic lines produced have been generated utilizing the SV40 oncogene and either the tyrosinase-promoter or the tyrosine-related protein promoter.75-80 Careful analysis suggests that the tumors begin in the neonatal period as a peripapillary multilayered proliferation of retinal pigment epithelial (RPE) cells. The early tumor cells are characterized by a spindle shape, abundant cytoplasm, round nuclei with uniform staining, and fine melanin pigment.78 Retinal, choroidal, and optic nerve invasion occurs by 6 to 10 weeks. By the end of this time the cells develop an appearance similar to human choroidal melanoma cells, including increased basophilia, nuclear and cytoplasmic polymorphism, prominent nucleoli, and abundant mitoses. With age, there is continued tumor growth with retinal detachment and extrascleral extension.78 However, the exact histogenesis of these pigmented tumors is confusing. While they appear to arise from RPE cells in some cases, they have many similarities to human uveal melanoma including morphology, tendency to metastasize, and expression of S-100 and HMB-45 antigens.77,78 In some instances the primary tumors apparently originate from the RPE;78,81 in others, from the RPE-choroid interface;77 and in still others, choroidal tumor formation occurs in the presence of normal RPE.80 Considering the neuroepithelial origin of RPE and the neural crest origin of choroidal melanocytes (see Chapter 3), this may be a nontrivial issue when studying molecular mechanisms of tumorigenesis. There are several potential explanations. The transgenic mice were produced on a variety of genetic backgrounds, and modifier genes may account for differences in the morphological phenotype. The presence of cutaneous melanomas in some studies may support this hypothesis,75,76 although in other studies the eye was the only tissue affected by tumors.78 Differences in construction of the transgene and use of different promoters might have a similar effect. A second possibility is that the reported differences were the result of dissimilar ages of the mice examined and consequent variability in the amount of tumor development. Apparent differences in site of tumor development could relate to the level at which sections were cut (see Chapter 13). However, the most likely explanation for the differences in transgene expression is that the RPE is more sensitive than the relatively inactive uveal melanocytes that also fail to express SV40 T antigen and that the RPE is the true source of origin.78,81 Further evidence supporting this hypothesis is demonstrated when the kit oncogene allele, KitW-v

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FIGURE 9.8 Retinoblastoma and tumor mimics. A. Retinoblastoma from a human eye with numerous rosettes (arrow). These consist of small cells arranged around a central lumen with eosinophilic cytoplasm on the luminal side. They resemble early photoreceptors. Reports of experimental retinoblastoma in mice document similar findings (see text). Original magnification × 400. B. Retinoblastoma often outgows its blood supply and becomes necrotic (*) or develops mineralization (arrow). C. Developmental thinning of the corneoscleral junction leads to a staphyloma (*) and may be lined by iris and ciliary body. This will appear as a dark spot on the surface, possibly suggesting a neoplasm. D. In a mouse homozygous for the ipd mutation, a dense accumulation of pigment-laden cells fills the anterior chamber, trabecular meshwork, aqueous collector channels, and adjacent sclera (arrow), also mimicking a neoplasm. Original magnification × 100. E. Under stress, the RPE can proliferate and also undergo metaplasia to form fibrous tissue (arrows). An extensive proliferation such as this can mimic a neoplasm. F. In addition to the anterior segment pigment dispersion and degeneration typical of DBA/2J mice, the choroid (arrow) often becomes depigmented in older mice. Original magnification × 200.

was introduced into the original Sv40-tyrosinase line.82 The KitW-v oncogene effectively ablated choroidal melanocytes while leaving the retinal pigment epithelium intact. In these mice, neoplasms clearly arose from the RPE. Although they were well differentiated, there was invasion of the optic nerve as well as more distant metastatic disease.76

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A related group of pigmented tumors that arise from the RPE has been produced in mice in which the ret oncogene was placed under the regulation of the mouse metallothionein promoter-enhancer (ocular and cutaneous pigmented tumors)83 or to the tyrosinase-related protein (Tyrp1) promoter. Both benign84 and malignant85 RPE tumors were observed in these transgenic mice. In mice with the Tyrp1 promoter, there was a high incidence of microphthalmia, perhaps partly related to the C57BL background of these mice.7 This further illustrates phenotypic variation with specific experimental conditions. There are several ocular histopathological findings that can clinically mimic pigmented neoplasms. The corneoscleral junction may become extremely thin and bulge externally after healing of a corneal ulcer. Peripheral corneal thinning is also a feature of some developmental abnormalities (see Chapter 8). In both instances, pigmented iris or ciliary body may line the bulging tissue, giving the appearance of an elevated pigmented tumor. This can usually be identified by shining a light through the pupil, which produces transillumination in the region of thinning (see Figure 9.8). Mice that develop pigment dispersion glaucoma (e.g., DBA/2J and AKXD-28/Ty86,87 may accumulate large numbers of pigmented cells in the anterior chamber, trabecular meshwork, and overlying sclera that can mimic a pigmented neoplasm. In the posterior segment, the retinal pigment epithelium retains the potential of proliferation. Especially after prolonged retinal detachment, large, elevated, multilayered plaques of pigment epithelium may suggest a neoplasm (see Figure 9.8).

CHOROIDAL DEGENERATION Degenerative changes in the choroid of mice are nearly always secondary to retinal diseases such as gyrate atrophy or other forms of retinal degeneration. Bruch’s membrane (derived partially from the choroid) and the choriocapillaris are most often affected. Diseases that fall in these categories are discussed in Chapter 10. More widespread choroidal degeneration occurs in aging DBA/2J mice. In older mice that have developed advanced secondary angle closure glaucoma, there is severe loss of pigment from the iris and ciliary body.86,88 Although not evident on ophthalmoscopic examination, there is marked transillumination of the posterior aspect of the enucleated eye, associated with extensive disappearance of choroidal pigment (see Figure 9.8).

LENS Formation of the lens begins around E10 and is controlled by complex molecular mechanisms (see Chapter 3).89-93 The lens is a simple structure consisting of the capsule, epithelium, and lens fibers (both cortex and nucleus). Its relative isolation from the systemic circulation and immune system limits the range of its reactions. Whereas trauma and specific forms of lens inflammation occur in human eyes,33 they are extremely rare in mouse eyes. Cataracts are the principal clinical and morphological phenotype observed in mouse lens. In its broadest definition, a cataract is an opacity in any part of the lens, although there may be considerable variation in morphology. Since many cataracts are visible without special equipment, they have been described since the earliest days of mouse genetics. The number of specific mouse cataract mutations has grown from 18 in 198294 to 62 in 199795 to well over 100 by 2001. Major reasons for the interest in cataracts are the frequent association of cataracts with other ocular and systemic developmental abnormalities, the discovery that genes that control lens formation also affect other ocular structures, and the fact that cataracts are a major cause of visual impairment in humans.7,25,96-98

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CATARACT TYPES Although most inherited cataracts in inbred mice are developmental in nature, age-related cataracts have also been reported.95,99-102 Some developmental cataracts demonstrate clinical progression during the first few months of life. A cataract may be first observed as a focal opacity that within a few weeks involves the entire lens cortex. Subtle clinical progression, especially in the lens nucleus, is best monitored with slit lamp examination. For example, the nucleus of a normal mouse lens is often difficult to identify, even with a slit lamp. When a nuclear cataract first appears, the lens nucleus initially becomes more distinct from the cortex before any opacification develops (Figure 9.9).

FIGURE 9.9 Cataract clinical phenotypes. A. Punctate cortical cataracts (arrow) are usually located in the lens cortex. B. Growth of the lens fibers from posterior to anterior results in the formation of X- or Y-shaped sutures that are often the site of sutural cataracts (arrow). C. In an eye with multiple developmental abnormalities there is a diffuse cataract that involves the entire lens. D. The center of the lens has a focal nuclear cataract. In addition, there are also diffuse cortical changes.

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CATARACT MORPHOLOGY General In a healthy mouse lens, the lens epithelium extends from the anterior pole of the lens to the equator, where it forms the lens bow (see Chapter 1). Posterior to the equator, the lens capsule is thinner than anteriorly and there is no lens epithelium. The initial morphological finding in many cataracts is posterior migration of the lens epithelium (Figure 9.10). A few epithelial nuclei beneath the posterior capsule are sufficient to diagnose cataract, even if the rest of the lens appears normal. It is important

FIGURE 9.10 Cataract. A. One of the earliest signs of cataract is posterior migration of the lens epithelium (arrow), which is ordinarily not present in this location. Original magnification × 200. B. When lens epithelial cells move posteriorly, they occasionally proliferate and produce abnormal lens proteins. These distended cells are referred to as bladder cells (arrow). Original magnification × 630. C. Congenital cataracts may appear before birth and frequently appear clinically and morphologically as vacuolation of the lens nucleus. Original magnification × 50. D. In another form of developmental cataract, a keratolenticular adhesion may occur (arrows). Concurrently, the anterior lens epithelium may proliferate, producing fibrous tissue. This particular example also shows foci of mineralization (arrowhead). Original magnification × 100. E. The presence of central nuclei (arrow) is also an indication of cataract. Original magnification × 100. F. The anterior lens capsule is thicker than normal in this eye with a combined nuclear and cortical cataract. A small patch of lens capsular material (arrow) indents the cortex. Original magnification × 250.

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to be aware of this early change when screening for morphological phenotypes. Especially in developmental cataracts, lens epithelial nuclei are often observed in the central lens. In cataracts of longer duration, displaced lens epithelial nuclei can produce malformed cortical material within their cytoplasm. This results in a coarsely granular cytoplasm that distends the cell, producing what is referred to as a “bladder cell” (Figure 9.10).95 Another early sign of cataract development is the presence of small cortical vacuoles. These are characteristically seen either beneath the anterior epithelium or in the lens bow area and can often be observed on slit lamp examination. Cataracts are usually advanced by the time they present as a white opacity in the pupil. Older mice with long-standing cataracts often develop focal mineralization in the degenerated cortex95 (see Figure 9.10). Capsular-Epithelial Because the lens capsule is a product of the lens epithelium, the two often react as a unit. Proliferation of the lens epithelium is frequently accompanied by thickening of the lens capsule and aberrant deposits of capsular material (Figure 9.11). Use of the “S” phase cell cycle marker bromodeoxyuridine (BrdU) demonstrates many lens epithelial cells that are premitotic (see Chapter 14A). On clinical examination, there is a white plaque of variable diameter and thickness directly beneath the anterior lens capsule that is termed an anterior subcapsular cataract (ASC). Because of the relatively undifferentiated status of lens epithelium, ASCs may undergo metaplasia and produce collagen.33,103 In older mice, cortical cataracts may develop. A typical example of an ASC is found in the translocation-induced circling mutation (Tim). Tim/+ mice develop an ASC after birth that is slowly progressive and is associated with abnormal head tossing and circling behavior. By 12 days of age the normal monolayer of lens epithelium beneath the anterior lens capsule has thickened to four to five cells. Adult mice develop a thickened ASC and lens capsule, lens epithelial proliferation, and fibrous metaplasia (see Figure 9.11). Some Tim/+ mice have a poorly differentiated hypoplastic iris and focal retinal dysplasia.104 Similar ASCs with epithelial proliferation and fibrous metaplasia occur in mice that express human platelet-derived growth factor-A (PDGFA) under the control of the αA crystallin promoter. These mice also develop thickening of the corneal stroma and delayed lid closure.105 Nuclear and Cortical The nucleus includes approximately 25% of the lens volume and is the earliest formed portion of the lens. In human eyes, slit lamp examination reveals a zone of discontinuity between nucleus and cortex that is easy to see. In mice, the nucleus is less easily distinguished, even with the use of higher magnification, except in the presence of a nuclear cataract. Since the nucleus is the oldest portion of the lens, the presence of a nuclear cataract suggests an event that occurred around E11 to E12, before the more external cortex develops. Typical nuclear cataracts occur in mutants with lens opacity 4 (Lop4),106 lens opacity 13 (lop13),107 and dominant cataract-2 (Cat2).108 Although cataracts may initially be limited to the nucleus, both cortex and capsule may become involved as the mouse ages.103 This tendency to progress accounts for the large number of spontaneous and targeted mutations that develop cortical and diffuse cataracts involving the entire lens (reviewed in Smith et al.95) (Figure 9.12). Cortical Liquefaction The lens cortex and nucleus of a longstanding cataract in human eyes may become completely liquefied, a condition known as hypermaturity.33 Although not previously described in mice, a very similar phenotype has been reported in the cataract and curly whiskers (Ccw) mutation.109 Although the lens developed normally in Ccw mice, cortical vacuoles were identified as early as P10. By 4 to 6 weeks of age, the entire cortex liquefies and by 14 weeks, only a vestige of the lens cortex remains,

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FIGURE 9.11 Anterior subcapsular cataracts. A. The lens epithelium may proliferate, forming multiple layers as well as focal capsular excrescences (arrow). A remnant of the anterior tunica vasculosa lentis is present (arrowhead). Original magnification × 400. B. This typical anterior subcapsular cataract is located at the anterior pole of the lens and consists primarily of aberrant deposits of lens capsular material. Original magnification × 250. C. The lens capsule shows marked thickening and the lens epithelial cells are growing in a fibroblastic pattern (arrow). D. A broad anterior subcapsular cataract contains equal parts of capsular and epithelial proliferation. Original magnification × 630. E. The lens cortex has disappeared and only the nucleus (N) remains. The folded lens capsule indicates the loss of subtance within the lens. The lens epithelium has undergone metaplasia to form a dense mat of fibroblast-like cells. Original magnification × 200. F. The vigorous uptake of the cell cycle marker (arrows), BrdU, indicates that the lens epithelium is undergoing active proliferation. Original magnification × 400.

surrounded by capsule. Presumably, the degenerated protein leaks out of the capsule, as often happens in hypermature cataract in human eyes. No anterior or posterior chamber cellular reaction was demonstrated in these mice, although this occurs frequently in human eyes.33,109 Similar findings were identified in an old FVB/N mouse (see Figure 9.12).

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FIGURE 9.12 Cataract types. A. In this (FVB/NJ-CAST/Ei) F1 cross, there is a prominent nuclear cataract (*). Original magnification × 100. B. The cataract in mice carrying the Lop 10 mutation demonstrates extensive vacuole formation (arrow) in the anterior lens cortex. Original magnification × 200. C. Both nuclear and cortical degeneration produce a diffuse cataract. Original magnification × 40. D. Cortical vacuoles, central nuclei, and swelling of lens fibers are seen in a higher magnification of C. Original magnification × 200. E. In longstanding cataracts, the lens proteins may become liquefied (arrow). An anterior subcapsular cataract (arrowhead) is also present. Original magnification × 100. F. In this lens, all of the lens cortex has become liquefied and the lens nucleus (*) has settled close to the posterior lens capsule. An anterior subcapsular cataract (arrow) appears in a lateral position because the lens capsule has partially collapsed as a result of the cortical changes. Original magnification × 100.

Lens Extrusion The posterior lens capsule is normally thinner than the anterior capsule (see Chapter 1). The thin posterior capsule may explain why this is usually the site of capsular rupture in lens extrusion cataracts. Some example of mutations associated with spontaneous capsular rupture include ectopic (ec),110 lens rupture (lr),111,112 lens opacity 10 (Lop10),113 and lens opacity 12 (Lop12, an allele of gamma D crystallin),103 as well as a variety of targeted mutations (reviewed in Smith et al.95). The

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common denominator in all lens rupture cataracts is a break in the posterior lens capsule that may happen close to birth or several weeks postpartum. Clinically, this occurrence is often marked by a sudden increase in cataract density. Morphologically, the ruptured capsule usually forms a coil because of its inherent elasticity. Lens cortical material spills into the vitreous and may float into the anterior chamber or become loosely attached to the surface of the retina (Figure 9.13). Unlike traumatic lens rupture in human eyes,33 the widespread dissemination of lens cortex in mouse eyes does

FIGURE 9.13 Lens extrusion cataracts. A. In this FVB/NJ mouse, the posterior lens capsule has ruptured (arrows) and lens cortex has extruded into the vitreous (V). There is prominent central migration of lens epithelium (arrowhead). Original magnification × 250. B. In an RBF/DnJ mouse, lens cortex is free in the vitreous (*) and lying on the retinal (R) surface, the latter inducing some retinal folding. Original magnification × 100. C. Shortly after capsular rupture, a large portion of the lens cortex fills most of the vitreous. The site of the capsular rupture is indicated by the arrow. Original magnification × 40. D. At higher magnification, the elasticity of the ruptured lens capsule has caused it to coil (arrow) at the point of lens extrusion. Original magnification × 630. E. The lens nucleus has extruded, leaving most of the lens cortex within the capsule. Original magnification × 40. F. Although lens extrusion usually does not produce inflammation, an inflammatory response does occur in Lop 12 mice. Here lens cortex is surrounded by reactive fibrous tissue in the anterior vitreous (arrow). Free cortical material is also present (arrowhead). Original magnification × 400.

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not stimulate acute or chronic inflammatory responses. The only reported exception to this is in lens opacity 12 (CrygdLop12/+) mice that develop a fibrovascular chronic inflammatory response in the vitreous (see Figure 9.13). Since the reaction is centered on extruded cortical fragments, it is reasonable to conclude that there is a direct relationship.103 Neoplasia In cataracts with lens epithelial proliferation, the response is cytologically benign. In fact, spontaneous neoplasms of the lens epithelium have not been reported in vertebrates. However, tumors do occur in genetically engineered mice when oncogenes or portions of tumor virus genomes (SV40 and papillomavirus type 16) are linked to lens crystallin promoters.114-117 Lens epithelial tumors confined within the lens capsule116 and invading other ocular structures including the orbit117 have been reported.

CATARACTS IN OCULAR AND SYSTEMIC DISEASE Development of the eye is a complex process that requires a host of molecular mechanisms controlled by multiple genes that affect multiple ocular tissues.89,96,118-120 Genes with mutations that cause ocular abnormalities in addition to cataracts include Pax6,97,98,121 Mitf,28 mammalian hairy and enhancer of split (Hes1),122 extra toes (Gli3Xt),123 and ocular retardation (Chx10or-J).124 Another unusual variant is the aphakia allele of paired-like homeodomain transcription factor 3 (Pitx3ak) in which lens development begins normally and then ceases with gradual disappearance of the lens primordia.125 Aphakia associated with microphthalmia have been reported in eye lens aplasia (elap)126 and also occurs in some forms of microphthalmia. The central corneal opacities, corneal thickening, retinal dysplasia, iris, and ciliary body malformations associated with these mutations were discussed earlier in Chapters 3, 4, and 8. Many different extraocular manifestations including craniofacial, skeletal, and central nervous system abnormalities are also associated with these mutations. Clearly, there are complex gene interactions that cause not only cataracts, but many other developmental difficulties as well.

VITREOUS The vitreous in mice, compared to human eyes, occupies a small portion of the ocular cavity.127 Since the vitreous body is normally acellular, consisting of highly dispersed collagen and hyaluronic acid, there is little to see in routine histological preparations. Between E10 and E11 (see Chapter 3) the hyaloid artery enters the eye by way of the choroidal fissure. The transient anastamosing network of intraocular vessels seen during development includes the hyaloid artery and its branches (the vasa hyaloidea propria) and the anterior and posterior tunica vasculosa lentis. The vessels located in the space between retina and lens are referred to as the primary vitreous. The hyaloid vascular system normally persists as long as P30 in mice. Hyaluronic acid and collagen appear around E14. Shortly after birth, involution of the hyaloid vascular system begins with apoptosis of the vascular endothelial cells, a process closely associated with macrophage migration into the vitreous.128-130

DEVELOPMENTAL ABNORMALITIES Vitreous Agenesis Microphthalmic eyes frequently contain little or no vitreous, particularly in those eyes with severe retinal abnormalities. It is possible for vitreous development to begin, but never be completed. This occurs in the anterior pyramidal cataract allele of dominant cataract 4 mutation, Cat4Apcat1-1. The early hyaloid vascular system can be identified in both Cat4 +/- and Cat4 -/- mice at E14. However, in

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homozygotes, hyaloid vessels are less developed and there is no evidence of a tunica vasculosa lentis that is clearly seen in wild-type littermates. Heterozygous adults are characterized by cataract and persistent attachment of the lens stalk to the cornea, but the vitreous is normal. The early hyaloid vascular system atrophies by birth in homozygotes and adult vitreous cannot be identified in the microphthalmic eyes. In addition, these mice are characterized by anterior segment dysgenesis, diffuse cataract, and retinal dysplasia.120 In addition to the corneal abnormalities discussed in Chapter 8,131 mice with two different mutations in the Col2a1 gene develop significant alterations in vitreous structure. The density of vitreous filaments is reduced, the individual vitreous filaments are thickened, and empty cavities develop within the vitreous substance. In addition, the overall volume of the vitreous chamber appears reduced (Figure 9.14).131

FIGURE 9.14 Vitreous. A and D. C57BL/6 × DBA/2J mixed background normal control, E18.5. The vitreous cavity (V) is of normal size and the vitreous network of fine-diameter fibrils is dense and in intimate contact with the lens (L), retina (R), and internal limiting membrane (ILM). B and E are Del3 transgenic line mice. C and F are Gly-85-1 transgenic mice. In both transgenic lines, the vitreous filaments are sparse, thickened, and empty cavities are present within the vitreous. The overall size of the vitreous cavity is reduced. A to C original magnification × 200; D to F, × 400. (From Ihanamaki, T., Eur. J. Ophthalmol., 6:427–435, 1996. With permission of Wichting Editore.)

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Persistent Hyperplastic Primary Vitreous The fetal vitreous vasculature, including the hyaloid artery, begins to disappear shortly after birth and is normally gone by 30 days of age in mice (Figure 9.15). Failure to complete this process results in various types of developmental abnormalities. The most common isolated anomaly is either small remnants of the hyaloid artery, usually extending a short distance from the optic nerve into the vitreous, or persistence of a few small vessels adjacent to the lens. Several mouse mutations have been described in which larger numbers of vessels persist, often accompanied by dense fibrous tissue proliferation in the

FIGURE 9.15 Vitreous. A. Anterior hyaloid vessels (arrow) in a 10-day-old C57BL/6J mouse. These are normally seen up to P30. When sections are cut through the lens bow, it is normal to see more central nuclei (arrowhead). Original magnification × 200. B. Small remnants of the hyaloid artery (arrow) may occur without other ocular abnormalities. C. This 7-day-old C57BL/6 mouse has more anterior and posterior hyaloid vessels than normal for this age. In addition, there is a nodule of pigmented tissue (arrowhead) posterior to the lens. Original magnification × 100. D. At higher magnification, the collection of pigmented cells (arrowhead) is closely associated with the hyaloid vessels, in a pattern similar to persistent hyperplastic primary vitreous. Original magnification × 400. E. In a p53-null mouse, the retina is pulled toward the lens by partially pigmented tissue. Original magnification × 100. F. At higher magnification, the pigmented tissue is tightly attached to the posterior lens capsule and to nonpigmented tissue adherent to the retinal surface. Original magnification × 400.

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vitreous or on the posterior surface of the lens. This is referred to as persistent hyperplastic primary vitreous (PHPV). PHPV is a common congenital abnormality in human eyes. Initially considered a single entity, it has subsequently been described in several different human syndromes (reviewed in Goldberg132). Both genetically and chemically induced forms of PHPV occur in mice. PHPV in p53-Null Mice The tumor suppressor gene Trp53 (hereafter, p53) was initially known for its important functions in tumor biology,133 especially relating to apoptosis and angiogenesis.134,135 Subsequently, Trp53 has been shown to be important in normal cranial and central nervous system development as some p53-null mice develop exencephaly.136 At certain stages of lens development, apoptosis is p53 dependent.137 Severe ocular abnormalities occur in p53-null mice on the C57BL/6J background. Abnormal vessels extend from the peripapillary area throughout the vitreous and become incorporated in retrolental fibrovascular membranes. Some of the retrolental membranes are pigmented, perhaps due to migration of mobilized RPE cells. In some mice, the retrolental membranes are associated with cataracts and early extrusion of the lens cortex. These findings resemble the human PHPV phenotype. In addition, mice show unilateral or bilateral hypoplasia of the optic nerves with diminution in number of nerve fibers, disorganized pial septae, and evidence of focal axonal degeneration. A corresponding focal retinal ganglion cell loss is also demonstrated. In contrast, p53-null mice on a 129/Sv background had only small hyaloid vessel remnants and no PHPV. The different expression on different genetic backgrounds (see Chapters 4 through 6) suggests the presence of a modifier gene that affects the disease phenotype (see Figure 9.15).138 PHPV in Norrie’s Disease In humans, Norrie’s disease is a sex-linked recessive syndrome characterized by fibrovascular preretinal proliferation, secondary retinal detachment, abnormal vitreous vessels, and the presence of a retrolental mass. In addition, there is mental retardation, progressive hearing loss, and epilepsy in many patients.33,139 The Norrie’s disease homologue (Ndph), which is mutated in the mouse model of Norrie’s disease, is expressed in retina, brain, and olfactory bulb of young mice (Figure 9.16).140 Hemizygous mice with a replacement mutation of exon 2 of the Ndph gene develop retrolental fibrovascular membranes and fibrous vitreous opacities. In addition there is disorganization of the ganglion cell layer with dislocated ganglion cells in the inner plexiform layer. Older mice also develop photoreceptor loss and retinoschisis (a splitting of the retinal layers).139-142 PHPV with Ectopic Gene Expression The complexity of the processes that govern ocular development is illustrated by the effects of two totally different genes that produce multiple ocular abnormalities that include PHPV. Factor VIII–associated gene B (F8B) is a small gene nested within the human coagulation factor VIII gene.143 When this human gene is inserted into mice there is severe growth retardation, microcephaly, and fibrovascular pigmented retrolental tissue that resembles PHPV. These mice, which were produced on a C57BL/6 background, also have microphthalmia, retinal dysplasia, and characteristic B6 corneolenticular adhesions (see Chapter 8). A similar range of abnormalities was found in mice genetically engineered to overexpress the mouse INF-γ gene in the lens.144,145 Some of these defects could also be explained as effects related to the site of transgene insertion. PHPV Induced by Retinoic Acid and Its Receptors Vitamin A (retinol) and its derivatives play a critical role in normal development and in maintenance of many tissues in the adult. The nuclear retinoic acid receptors are regulatory factors that control retinoic acid metabolism. Deficiency or excess of vitamin A is known to produce developmental

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FIGURE 9.16 Mouse homologue of Norrie’s disease. A. A 148-day-old control mouse. The retina is normally developed and blood vessels (arrows) are present throughout the inner retina. B. In a mouse with a model of Norrie’s disease, vessels are present only at the nerve fiber–ganglion cell layer level (arrow) and the photoreceptors are not full developed. C. In a different mouse, vessels appear located at the vitreoretinal interface, suggesting extension into the peripheral vitreous. There is severe maldevelopment of the photoreceptor layer. D to G. Adult mice with the Norrie’s disease model. D. A vessel from the inner plexiform layer has a collapsed lumen. E. On the inner retinal surface, vessels are not covered by Muller cells, only by the internal limiting membrane (arrows). The endothelium is attenuated. F. In some places, vessels penetrate the internal limiting membrane (ILM, arrowhead). The vitreal and retinal vessels are fenestrated (arrows), although normal retinal vessels are not. G. Larger vessel with pericytes in the vitreous adjacent to the internal limiting membrane (ILM, arrowhead), surrounded by fibrous material. (From Lutjen-Drecoll, E., Invest. Ophthalmol. Vis. Sci., 39:2450–2457, 1998. With permission.)

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defects that include cardiac, craniofacial, and skeletal abnormalities (reviewed in Ghyselinck et al.146). Double null mutants involving several isoforms of retinoic acid receptors develop ocular abnormalities that include PHPV, ocular colobomas, and anterior segment dysgenesis in addition to the systemic defects.3,5,146 It is of interest that an excess of retinoic acid is capable of producing similar effects. When exogenous retinoic acid is administered to pregnant mice prior to day 12, there is striking fibrovascular proliferation in the vitreous that surrounds the posterior aspect of the lens.8,147

VITREOUS HEMORRHAGE Vitreous hemorrhage in mice usually is related to underlying ocular disease. The presence of abnormal vessels in the vitreous either from persistent fetal vasculature or arising from the retina (see Chapter 10) may rarely be associated with bleeding. Older mice are more likely to develop a vitreous hemorrhage and some strains, such as DBA/2 or PL/J, are particularly prone to spontaneous intraocular hemorrhage. Blood can remain unchanged in the vitreous for long periods of time, but eventually breaks down with deposition of free hemosiderin in the vitreous or contained within mobilized macrophages. Whole blood cells may also be engulfed by macrophages. Vitreous hemorrhage is a potential complication when blood is drawn from the orbital venous sinus, although intraorbital hemorrhage is more likely (Figure 9.17).

VITREOUS INFLAMMATION When inflammation does occur in the vitreous, it is normally secondary to an exogenous stimulus. For example, a severe corneal ulcer may allow intraocular invasion of bacteria, which can lead to widespread intraocular inflammation (endophthalmitis) that includes the vitreous. A more severe level of inflammation can lead to involvement of uvea, sclera, and orbit (panophthalmitis). The inflammation associated with experimental autoimmune uveitis and experimental herpes simplex infections was discussed earlier in this chapter.

FIGURE 9.17 A. In an old vitreous hemorrhage (*), brown pigment within the hemorrhage is probably hemosiderin (arrow). B. In a large subretinal hemorrhage, cells that are either macrophages or free RPE cells have engulfed erythrocytes, a process termed erythrophagocytosis. Original magnification × 630.

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82. Mintz, B. and Klein-Szanto, A., Malignancy of eye melanomas originating in the retinal pigment epithelium of transgenic mice after genetic ablation of choroidal melanocytes, Proc. Natl. Acad. Sci. U.S.A., 89:11421, 1992. 83. Iwamoto, T. et al., Aberrent melanogenesis and melanocytic tumor development in transgenic mice that carry a metallothionein/ret fusion gene, EMBO J., 10:3167, 1991. 84. Schmidt, A. et al., Ectopic expression of RET results in microphthalmia and tumors in the retinal pigment epithelium, Int. J. Cancer, 80:600, 1999. 85. Beerman, F., Hunziker, A., and Foletti, A., Transgenic mouse models for tumors of melanocytes and retinal pigment epithelium, Pigment Cell Res., 12:71, 1999. 86. John, S.W.M.J. et al., Essential iris atrophy, pigment dispersion and glaucoma in DBA/2J mice, Invest. Ophthalmol. Vis. Sci., 39:951, 1998. 87. Anderson, M.G. et al., Genetic modifications of glaucoma associated phenotypes between AKXD28/Ty and DBA/2J mice, BMC Genet., 2:1, 2001. 88. Chang, B. et al., Interacting loci cause severe iris atrophy and glaucoma in DBA/2J mice, Nat. Genet., 21:405, 1999. 89. Collinson, J.M., Hill, R.E., and West, J.D., Different roles for Pax6 in the optic vesicle and facial epithelium mediate early morphogenesis of the murine eye, Development, 127:945, 2000. 90. Desplan, C., Eye development: governed by a dictator or a junta? Cell, 91:861, 1997. 91. Fini, M.E., Strissel, K.J., and West-Mays, J.A., Perspectives on eye development, Dev. Genet., 20:175, 1997. 92. van Raamsdonk, C.D. and Tilghman, S.M., Dosage requirements and allelic expression of PAX6 during lens placode formation, Development, 127:5439, 2000. 93. Schwarz, M. et al., Spatial specification of mammalian eye territories by reciprocal transcriptional repression of Pax2 and Pax6, Development, 127:4325, 2000. 94. Robison, W.G., Kuwabara, T., and Zwaan, J., Eye research, in Foster, H.L., Small, J.D., and Fox, J.G., Eds., The Mouse in Biomedical Research, Academic Press, New York, 1982, 80. 95. Smith, R.S., Sundberg, J.P., and Linder, C.C., Mouse mutations as models for studying cataracts, Pathobiology, 65:146, 1997. 96. Graw, J., Genetic aspects of embryonic eye development in vertebrates, Dev. Genet., 18:181, 1996. 97. Schedl, A. et al., Influence of PAX6 gene dosage on development: overexpression causes severe eye abnormalities, Cell, 86:71, 1996. 98. Quinn, J.C., West, J.D., and Hill, R.E., Multiple functions for Pax6 in mouse eye and nasal development, Genes Dev., 10:435, 1996. 99. Kuck, J.F.R., Late onset hereditary cataract of the Emory mouse. A model for human senile cataract, Exp. Eye Res., 50:659, 1990. 100. Hosokawa, M. et al., Persistent hyaloid vascular system in age-related cataract in a SAM strain of mouse, Exp. Eye Res., 57:427, 1993. 101. Ashida, Y., Takeda, T., and Hosokawa, M., Protein alterations in age-related cataract associated with a persistent hyaloid vascular system in senescence-accelerated mouse (SAM), Exp. Eye Res., 59:467, 1994. 102. Li, Y., Yan, Q., and Wolf, N.S., Long-term caloric restriction delays age-related decline in proliferation capacity of murine lens epithelial cells in vitro and in vivo, Invest. Ophthalmol. Vis. Sci., 38:100, 1997. 103. Smith, R.S. et al., Lop12, a mutation in mouse Crygd causing lens opacity similar to human Coppock cataract, Genomics, 63:314, 2000. 104. Smith, R.S. et al., Cataract and neurological abnormalities in partial trisomy, Invest. Ophthalmol. Vis. Sci., 38:S585, 1997. 105. Reneker, L.W. and Overbeek, P.A., Lens-specific expression of PDGF-A in transgenic mice results in retinal astrocytic hamartomas, Invest. Ophthalmol. Vis. Sci., 37:2455, 1996. 106. West, J.D. and Fisher, G., Further experience of the mouse dominant cataract mutation test from an experiment with ethylnitrosourea, Mutant Res., 1986 164:127. 107. Varnum, D., Nuclear cataract (nuc), Mouse News Lett., 64:59, 1981. 108. Graw, J. et al., Characterization of Cat-2, a radiation-induced dominant cataract mutation in mice, Invest. Ophthalmol. Vis. Sci., 31:1353, 1990.

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109. Lyon, M.F. et al., Further genetic analysis of two autosomal dominant mouse eye defects, Ccw and Pax6coop, Mol. Vis., 6:199, 2000. 110. Beasley, A.B., Inheritance and development of a lens abnormality in the mouse, J. Morphol., 112:1, 1963. 111. Fraser, F.C. and Herer, M.L., The inheritance and expression of the “lens rupture” gene in the house mouse, J. Hered., 41:3, 1950. 112. Fraser, F.C. and Herer, M.L., Lens rupture, a new recessive gene in the house mouse, J. Hered., 39:149, 1948. 113. Runge, P.E. et al., Autosomal dominant mouse cataract (Lop-10): consistent differences of expression in heterozygotes, Invest. Ophthalmol. Vis. Sci., 33:3202, 1992. 114. Griep, A.E. et al., Tumorigenicity by human papillomavirus type 16 E6 and E7 in transgenic mice correlates with alterations in epithelial cell growth and differentiation, J. Virol., 67:1373, 1993. 115. Gotz, W. et al., Eye pathology in transgenic mice carrying a MSV-SV 40 large T-construct, Exp. Eye Res., 52:41, 1991. 116. Pichel, J.G., Lakso, M., and Westphal, H., Timing of SV40 oncogene activation by site-specific recombination determines subsequent tumor progression during murine lens development, Oncogene, 8:3333, 1993. 117. Mahon, K.A. et al., Oncogenesis of the lens in transgenic mice, Science, 235:1622, 1987. 118. Oliver, G. and Gruss, P., Current views on eye development, Trends Neurosci., 20:415, 1997. 119. Dahl, E., Koseki, H., and Balling, R., Pax genes and organogenesis, Bioessays, 19:755, 1997. 120. Grimes, P.A. et al., Abnormal eye development associated with Cat4a, a dominant mouse cataract mutation on chromosome 8, Invest. Opthalmol. Sci., 39:1863, 1998. 121. Kaufman, M.H., Chang, H.H., and Shaw, J.P., Craniofacial abnormalities in homozygous small eye (Sey/Sey) embryos, J. Anat., 186:607, 1995. 122. Tomita, K. et al., Mammalian hairy and enhancer of split homolog-1 regulates differentiation of retinal neurons and is essential for eye morphogenesis, Neuron, 16:723, 1996. 123. Hui, C. et al., Expression of three mouse homologs of the Drosophila segment polarity gene cubitus interruptus, Gli, Gli-2, and Gli-3, in ectoderm- and mesoderm-derived tissues suggests multiple roles during postimplantation development, Dev. Biol., 162:402, 1994. 124. Burmeister, M. et al., Ocular retardation mouse caused by Chx10 homeobox null allele: impaired retinal progenitor proliferation and bipolar cell differentiation, Nat. Genet., 12:376, 1996. 125. Grimm, C. et al., Aphakia (ak), a mouse mutation affecting early eye development: fine mapping, consideration of candidate genes and altered Pax6 and Six3 gene expression pattern, Dev. Genet., 23:299, 1998. 126. Aso, S.S.H. and Noda, S., Lens aplasia: a new mutation producing lens abnormality in the mouse, Lab. Animal Sci., 45:41, 1995. 127. Hogan, M.J., Alvarado, J.A., and Weddell, J.E., Histology of the Human Eye, W.B. Saunders, Philadelphia, 1971. 128. Lang, R. et al., Apoptosis during macrophage-dependent ocular tissue remodeling, Development, 120:3395, 1994. 129. Lang, R.A. and Bishop, J.M., Macrophages are required for cell death and tissue remodeling in the developing mouse eye, Cell, 74:453, 1993. 130. Ito, M. and Yoshioka, M., Regression of the hyaloid vessels and pupillary membrane of the mouse, Anat. Embryol. Berl., 200:403, 1999. 131. Ihanamaki, T. et al., Ocular abnormalities in transgenic mice harboring mutations in the type II collagen gene, Eur. J. Ophthalmol., 6:427, 1996. 132. Goldberg, M.F., Persistent fetal vasculature (PFV): an integrated interpretation of signs and symptoms associated with persistent hyperplastic primary vitreous, Am. J. Ophthalmol., 124:587, 1997. 133. Merlino, G., Regulatory imbalances in cell proliferation and cell death during oncogenesis in transgenic mice, Semin. Canc. Biol., 5:13, 1994. 134. Rak, J., Filmus, J., and Kerbel, R.S., Reciprocal paracrine interactions between tumor cells and endothelial cells: the “angiogenesis progression” hypothesis, Eur. J. Cancer, 32A:2438, 1996. 135. Stellmach, V. et al., Tumor suppressor genes and angiogenesis: the role of TP53 in fibroblasts, Eur. J. Cancer, 32A:2394, 1996.

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136. Sah, V.P. et al., A subset of p53-deficient embryos exhibit exencephaly, Nat. Genet., 10:175, 1995. 137. Pan, H. and Griep, A.E., Temporally distinct patterns of p53-dependent and p53-independent apoptosis during mouse lens development, Genes Dev., 9:157, 1995. 138. Ikeda, S. et al., Ocular abnormalities in C57BL/6 but not 129/Sv p53 deficient mice, Invest. Ophthalmol. Vis. Sci., 40:1874, 1999. 139. Berger, W., Molecular dissection of Norrie disease, Acta Anat., 162:95, 1998. 140. Berger, W. et al., An animal model for Norrie disease (ND): gene targeting of the mouse ND gene, Hum. Mol. Genet., 5:51, 1996. 141. Ruether, K. et al., Retinoschisis-like alterations in the mouse eye caused by gene targeting of the Norrie disease gene, Invest. Ophthalmol. Vis. Sci., 38:710, 1997. 142. Richter, M. et al., Retinal vasculature changes in Norrie disease mice, Invest. Ophthalmol. Vis. Sci., 39:2450, 1998. 143. Valleix, S. et al., Expression of human F8B, a gene nested with the coagulation factor VIII gene, produces multiple eye defects and developmental alterations in chimeric and transgenic mice, Hum. Mol. Genet., 8:1291, 1999. 144. Egwuagu, C.E. et al., Ectopic expression of gamma interferon in the eyes of transgenic mice induces ocular pathology and MHC Class II gene expression, Invest. Ophthalmol. Vis. Sci., 35:332, 1994. 145. Egwuagu, C.E. et al., γ Interferon expression disrupts lens and retinal differentiation in transgenic mice, Dev. Biol., 166:557, 1994. 146. Ghyselinck, N.B. et al., Role of the retinoic acid receptor beta (RARβ) during mouse development, Int. J. Dev. Biol., 41:425, 1997. 147. Ozeki, H. et al., Critical period for retinoic acid-induced developmental abnormalities of the vitreous in mouse fetuses, Exp. Eye Res., 68:223, 1999.

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Retina Richard S. Smith, Norman L. Hawes, Bo Chang, and Patsy M. Nishina

CONTENTS Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .195 Developmental Abnormalities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .196 Retinal Dysplasia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .196 Microphthalmia Alleles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .196 Retinal Vascular Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .198 Retinopathy of Prematurity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .198 Primary Photoreceptor Degenerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .198 Retinal Degeneration-1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .201 Retinal Degeneration-2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .201 Retinal Degeneration-3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .204 Retinal Degeneration-4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .204 Retinal Degeneration-5 and Tubby-Like Protein-1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .204 Retinal Degeneration-6 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .207 Retinal Degeneration-7 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .207 Purkinje Cell Degeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .208 Nervous . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .211 Neuronal Ceroid Lipofuscinoses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .211 Cone-Rod Homeobox Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .214 Arrestin—Oguchi’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .215 Rhodopsin Mutations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .216 Other Retinal Degenerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .216 Retinal Detachment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .216 Primary Diseases of the Retinal Pigment Epithelium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .218 Dominant Negative FGF Receptor-1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .218 Gyrate Atrophy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .218 Senescence-Accelerated Mouse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .218 Degenerative and Reactive Changes in the RPE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .220 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .221

INTRODUCTION Both retina and retinal pigment epithelium (RPE) arise from neuroepithelium. Their close relationship is emphasized by the capability of the RPE during development to be transdifferentiated into retina, and vice versa.1,2 The functional relationship between retina and RPE is maintained in adult life with the RPE forming a part of the blood–retinal barrier between the choriocapillaris and the outer retina.3,4 The RPE is responsible for generating the extracellular matrix that surrounds the outer segments of the photoreceptors. Perhaps the most critical role of the RPE is degradation of cast-off

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outer segments and regeneration of visual pigments.5-8 Many retinal and RPE diseases also involve Bruch’s membrane and the choriocapillaris (see Chapter 9).

DEVELOPMENTAL ABNORMALITIES RETINAL DYSPLASIA The term retinal dysplasia is used to describe a developmental abnormality in which tubular or rosette-like structures are found in the retina. It has been described in human eyes,9 as well as in other animals.10 Although originally thought to be a specific diagnosis (reviewed in Lahav and Albert9), it has been found in many different human11 and mouse diseases. In addition to genetic factors, maternal viral infections are associated with retinal dysplasia.9 Dysplasia has been reported in several different mouse gene knockouts including those of retinoic acid receptors,12 α-crystallin,13 p27Kip1 mice,14 and mice lacking p56lck.15 Retinal dysplasia also occurs in the mouse model of Norrie’s disease (see Chapter 9),16 in mice with a transgene involving elements of the Moloney murine sarcoma virus and the Simian virus 40 large T antigen,17 as a background lesion in inbred black mice,18 and in partial trisomy of mouse Chr 4 and 17.19 Morphology of the dysplastic retina is similar regardless of the molecular mechanism. It seems reasonable to conclude that retinal dysplasia is a nonspecific response to diverse stimuli that affect retinal differentiation during embryonic development. Retinal dysplasia may involve the entire retina, but is most commonly a focal abnormality (Figure 10.1). Differentiation of the tubules and rosettes is variable, from one to two layers of cells to multilaminar structures surrounding a lumen. The cells adjacent to the lumen may be ciliated, a characteristic of primitive neuroepithelium.

MICROPHTHALMIA ALLELES The microphthalmia (mi) mutant, originally described more than 60 years ago, is characterized by skeletal (osteopetrosis) and dental abnormalities, diminished pigmentation of hair and eyes, microphthalmia, and deafness (reviewed in Packer20). More than a dozen alleles, both dominant and recessive, have been described at the microphthalmia locus. The phenotype of these alleles often demonstrates variations from the original phenotype.21,22 These are caused by multiple mutations in the microphthalmia-associated transcription factor (Mitf), a member of the basic-helix-loop-helixleucine zipper family of transcription factors.21,23 MITF plays a critical role in the selective differentiation of neuroepithelium into neural retina or RPE early in development.24,25 Despite the interest in Mitf alleles, for the most part, the published illustrations of the ocular morphology leave many unanswered questions concerning the frequency, consistency, and extent of abnormalities. Because of the limited information in published reports, it is not possible to discuss the ocular morphological characteristics of each allelic mutation. The problem is further complicated by the fact that coat color of the background strain affects ocular findings.22 The most commonly reported abnormalities include microphthalmia, decrease in uveal pigmentation, proliferation of retinal pigment epithelium, retinal degeneration, and colobomas of the retina and optic nerve. Absence or severe decrease in uveal pigmentation occurs in both choroid and iris. This results in increased iris transillumination on slit lamp examination and enhanced transparency of the posterior globe in the prolapsed or enucleated eye. Iris stromal melanocytes are frequently absent and there appears to be less pigment than normal in the iris pigment epithelium in those allelic mutations with a pigmented coat.26-28 Relatively normal iris pigmentation may be present at the same time that choroidal pigment is virtually absent. In some alleles, such as Mitfmi-vit, choroidal pigmentation is present peripherally, but absent in the posterior globe (see Figure 10.1).20 This allele is also characterized by the presence of numerous melanophages in both choroid and iris that resemble those found

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FIGURE 10.1 Retinal dysplasia and microphthalmia. A. Multiple foci of dysplastic retina (arrow) are located primarily in the inner nuclear layer. Original magnification × 200. B. At higher power, the nuclei are polarized away from a “lumen.” The arrowhead indicates structures resembling photoreceptor inner segments and the linear structure (arrow) is similar to the external limiting membrane of the retina. This supports the view that dysplastic rosettes arise from misdirected primitive neuroblastic cells. Original magnification × 630. C. Comparing the nuclear appearance, this isolated rosette appears to arise from the outer nuclear layer. In this Bst mouse, there are persistent hyaloid vessels in the adult (arrow). Original magnification × 400. D. In this Bmp4 mouse, the entire outer nuclear layer consists of dysplastic rosettes. A recent vitreous hemorrhage (H) lies on the retinal surface. Original magnification × 200. E. Mice homozygous for some alleles of the microphthalmia gene (Mitf) fail to develop normal melanin pigmentation in the choroid (arrow). Original magnification × 630. F. In the same mouse as E, the iris stromal and epithelial pigmentation is normal. Original magnification × 630.

in older DBA/2J mice (see Chapter 8).28 For the most part, the appearance and pigmentation of the RPE is normal. However, in the Mitf Mi-wh allele there are focal areas of multilayered RPE.20 A late progressive photoreceptor degeneration was reported in both Mitfmi-vit and Mitfmi-di strains.21,27,29-31 At 3 to 4 months of age photoreceptor nuclei and outer segments degenerate. With increasing age these changes become more pronounced. The RPE of Mitfmi-vit mice contains less than

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half as many phagosomes as are found in normal C57BL/6 controls, suggesting that defective phagocytosis may play a role in the retinal degeneration.32 By 6 months of age, the entire photoreceptor layer disappears and the inner nuclear layer is in direct contact with the RPE (Figure 10.2). A cardinal finding in the original description of Mitfmi/mi mice was the presence of severe microphthalmia.33 However, some allelic mutations have eyes of normal size, or varying degrees of microphthalmia.21 The morphological findings in microphthalmic mice differ considerably from those found as a background lesion in inbred black mice in which nearly all parts of the eye are involved (see Chapter 9).18 In affected Mitf mutant mice, the entire eye is decreased in size to about 50% of normal, and the anterior segment of the eye may be abnormal. Microscopically, there are usually multiple foci of retinal dysplasia as well as a more diffuse irregularity of the retinal architecture, including coloboma of the optic nerve that contains prolapsed retina, RPE, and choroid (see Figure 10.2).20,33,34 The coloboma resembles those found in other mouse strains (see Chapter 11).

RETINAL VASCULAR DISEASE In human eyes the scope of retinal disease based on the retinal vascular system includes arterial and venous occlusion, hypertension, diabetes, and a host of other systemic diseases that affect the vascular system.11 Thus far, relatively few retinal vascular diseases have been described in mice. Choroidal neovascularization that extends into the subretinal space and produces retinal damage was discussed in Chapter 9. Because the choroidal circulation is critical for maintenance of the outer retina, choroidal vascular insufficiency as occurs in the mouse model of lupus erythematosis35 is probably responsible for damage to the RPE and photoreceptors (see Chapter 9).

RETINOPATHY OF PREMATURITY As the hyaloid vascular system involutes during the first 2 weeks of life in mice, the retinal vascular system develops and grows, beginning at the optic nerve and extending peripherally.36,37 In both mice and humans the developing retina is vulnerable to increased ambient oxygen levels. The growing vessels respond by undergoing severe vasoconstriction that persists as long as the oxygen level is high. Instead of resuming normal growth when atmospheric oxygen is restored, vessels grow through the internal limiting membrane of the retina into the vitreous. Tufts of vessels that develop on the surface of the retina and in the peripheral vitreous are prone to hemorrhage and leakage of plasma proteins. Blood and blood products induce vitreous fibrosis that leads to vitreoretinal traction and secondary retinal detachment.11 In human eyes, this condition is referred to as retrolental fibroplasia or retinopathy of prematurity (ROP). ROP at one time was a major cause of visual loss in infants and children. However, the incidence has decreased, as the source of the problem became evident. Because of the similarity between retinal vascular development in humans and mice, models of oxygen-induced retinopathy have been developed in the mouse. The most reliable seems to be exposure of 1-week-old mice to 75% oxygen for a 5-day period followed by a return to room air. ROP develops between P17 and P21 and is most prominent at the junction between vascularized and avascular retina in the midperiphery (Figure 10.3).37-39 Tufts of vessels grow through the internal limiting membrane into the adjacent vitreous. The full-blown human syndrome with vitreous fibrosis and retinal detachment has not been described in mice, although there are no reports of long-term clinical observations.

PRIMARY PHOTORECEPTOR DEGENERATIONS Many different genetically determined retinal diseases that end in cellular death and loss of photoreceptor function have been described in humans.11,40-44 Study of these diseases is frustrating, because

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FIGURE 10.2 Microphthalmia alleles. A. Mitfmi mouse 22 days of age. The cornea (C) is twice normal thickness and vascularized (arrowhead). The iris (I) is malformed and attached to the lens (L) by posterior synechias. There is a dense persistent pupillary membrane (arrow). Original magnification × 100. B. In the same mouse, the retina (R) is folded and incompletely differentiated. Original magnification × 100. C. In a central section through another Mitfmi mouse eye, the globe is about half normal size. The cataractous lens (L) nearly fills the eye, anterior segment structures cannot be identified, and the retina is folded. Original magnification × 50. D. A higher power shows the folded retina and a fragment of extraocular muscle (M) adjacent to the globe. The optic nerve should be in the space indicated by arrows, but has not formed. Original magnification × 100. E. In this 4-month-old Mitfvit/vit mouse, the posterior retina is normal. Original magnification × 400. F. In a 6-month-old Mitfvit/vit mouse, the ganglion cell (G) and inner nuclear layers (INL) are normal, but the photoreceptor nuclei have disappeared. Original magnification × 400.

they typically have a long course and availability of morphological material at early stages is rare. Molecular studies are difficult for the same reason and identification of causative genes requires large kindreds and diligent effort. When an apparently significant polymorphism is found,45 there is often conflicting evidence with regard to its relevance to a specific disease.46 Primary photoreceptor degenerations in mice have been known for many years. One of the earliest descriptions, of the

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FIGURE 10.3 Effects of hyperoxia in newborn mice. Animals exposed to 75% oxygen from P7-P12. A, C, and E. Whole isolated retinas, flat-mounted and observed under ultraviolet light after cardiac injection of fluorescein-conjugated dextran (molecular weight 2 × 106) (calibration bar: 400 µm). B, D, and F. Peripapillary retina stained with PAS and hematoxylin. A and B. Normal retina of an animal exposed only to ambient air. C and D. Retina at P12, after 5 days in hyperoxia. E and F. Retina at P17, 5 days after return to ambient air. aa = avascular area, c = penetrating capillary; ilm = internal limiting membrane, np = neovascularization process, nt = neovascular tuft. (Courtesy of Michel Lonchampt.)

“rodless” retina,47 was proved 70 years later by analysis that showed the DNA from the original slides to be the same as retinal degeneration-1 (Pde6brd1).48 Subsequently, several novel retinal degeneration models were described in mice and many genes have been mapped and cloned. Many of these models resemble human retinal degenerations and offer the opportunity to sequentially study clinical evolution, electrophysiology, and morphological changes. Insights into molecular mechanisms have been gained.41,43,44 Understanding of underlying mechanisms is one step away from the development

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of effective treatment. This possibility has been demonstrated in Pde6brd1 and retinal degeneration slow (Prph2) mice in which both structure and function can be restored by appropriate treatment.49,50 The Prph2 mice are of particular interest since some human outer retinal degenerations demonstrate similar molecular mechanisms.51,52 Many mouse mutations associated with retinal degeneration have been reported and the literature is extensive. However, a common denominator observed in all models is degeneration of the outer segments of the photoreceptors of rods, cones, or both. In some cases, other retinal layers and the retinal pigment epithelium are involved. The principal differences, in most cases, lie in the fundus appearance, the time of onset, the rate of progression, and the specific electrophysiological changes. As an introduction to the primary retinal degeneration models, Table 10.1 summarizes the major features of the different mutations and lists pertinent references.

RETINAL DEGENERATION-1 Keeler47 described the first retinal degeneration more than 75 years ago and called it the “rodless” retina (r). A morphologically similar mutation was called retinal degeneration-1 (rd1) and the identity of this mutation and Keeler’s original mice was established.48 Retinal degeneration-1 (Pde6brd1) is due to reduced activity of the β subunit of rod cGMP-phosphodiesterase.76,77 It is important for investigators evaluating eyes to be aware of Pde6brd1 and its associated morphological findings, as it is a frequent strain background disease (see Chapter 4). Strains that carry Pde6brd1 include BDP/J, BUB/BnJ, C3H/HeJ, C3H/HeJSx, C3H/HeOuJ, C3H/HeSnJ, C3HheB/FeJ, C3HfB/Bi, CBA/J, DA/HuSn, FL/1Re, FL/4Re, FVB/NJ, WB/ReJ KitW/+, NON/LtJ, P/J, PL/J, SB/Le, SJL/J, ST/bJ, SWR/J, WB/ReJ-+/+, WB/ReJ KitW/+, WC/ReJ-+/+, and WC/ReJ-MgfSl/+. This list is drawn from the JAX mouse product list (www.jax.org/jaxmice) and is by no means complete, because there are many variants of the listed strains. Investigators should be particularly cautious about strains with either C3H or FVB ancestry since they are likely to carry Pde6brd1. The frequent use of FVB strains to produce transgenic mice is a particular problem78 and has lead to incorrect interpretation of ocular morphology. Because many of the Pde6brd1 strains have albino coat colors that make clinical detection of the retinal disease more difficult, the retinal status should be confirmed by histology or by using specific markers for the mutation.77 At birth, the mouse retina is still undergoing development. Photoreceptor inner and outer segments are absent and do not begin to develop until P6 to P8 (postpartum day 6 to 8; see Chapter 3). Full growth of the outer segments is completed shortly before the eyes open at P14. By this time, mice homozygous for Pde6brd1 already exhibit signs of outer segment degeneration. The destruction proceeds rapidly and by P20 nearly all photoreceptor nuclei have disappeared (reviewed in LaVail and Mullen79). The initial impact is on the rods, while cones are preferentially spared. For example, at P17, only 2% of rods remain, while 75% of cones persist.53 The photoreceptors undergo apoptosis80 that is p53 independent.81 During the period of acute cell death, the RPE remains normal.79 The retinal vascular system develops normally in Pde6brd1 homozygotes, but begins to degenerate by P14, a process that continues with age. By 1 month of age, there is a 35% decrease in the number of vascular profiles.82 Clinical observation confirms the presence of vascular changes. By 2 months, retinal arterioles are pale and have no visible blood column.83 In older affected mice, the retinal vessels are often difficult to identify on fundus examination. This vascular abnormality likely contributes to the loss of retinal ganglion cells and nerve fibers observed in older Pde6brd1 mice (Figure 10.4).84

RETINAL DEGENERATION-2 This retinal degeneration model was originally called retinal degeneration slow (rds) because of its relatively late onset compared with Pde6brd1. The mutation (Prph2Rd2) responsible for the phenotype disrupts production of the rds/peripherin protein that is responsible for maintenance of photoreceptor

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TABLE 10.1 Mouse Models of Retinal Degeneration Mouse Mutation

Chromosome

Inheritance

Comments

Retinal degeneration-1, Pde6brd1

10

Recessive

Begins at P11 in ONL; some cones remain at 18 months

Retinal degeneration-2, Prph2Rd2

17

Retinal degeneration-3, rd-3

1

Retinal degeneration-4, Rd-4

Ref. 53

Semidominant Begins around P7 and progresses slowly

54, 55

Recessive

Initially normal; Onset 3 weeks in rods; cone remnants at 7 weeks

56

4

Dominant

ONL/OPL affected at P10; total loss by 6 weeks

57

Retinal degeneration-5 (tubby), tub

7

Recessive

Progressive ONL loss 2 weeks to 8 months

Retinal degeneration-6, rd-6

9

Recessive

Begins in ONL at P7 and progresses for 1 year; subretinal macrophages

Retinal degeneration-7, Nr2e3rd7

9

Recessive

Early retinal folding and late progressive ONL loss

61, 62

Tubby-like protein-1, Tulp1 –/–

17

Knockout

Rapidly progressive ONL loss starting at 3 weeks; inner segment vacuoles

63–65

Purkinje cell degeneration, pcd

13

Recessive

Slowly progressive ONL loss from P25; posterior retina more affected

66, 67

Nervous, nr

8

Recessive

Initial rapid ONL loss starting at P11–19; then slower progression through 17 months

68, 69

Motor neuron degeneration, mnd

8

Recessive

Progressive from 5 weeks to 6 months; peripheral retina worse initially

70, 71

Neuronal ceroid lipofuscinosis, nclf

9

Recessive

Similar to mnd, but slower

Cone-rod homeobox containing gene, Crx

7

Knockout

ONL normal at P14 with progressive loss through 6 months; absent outer segments

73, 74

Susceptibility to light damage (Arrb –/–)

7 (arrestin β 1) 11 (arrestin β 2)

Arrestin Knockout

ONL and outer segment loss with prolonged cyclic light exposure

22, 75

58, 59

60

72

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FIGURE 10.4 Retinal degeneration. A to D. Retinal degeneration-1 A. At age 24 days, there are coalescent patches of depigmentation (arrow). B. At age 61 days, depigmentation is more widespread and a geographic pattern has developed. The retinal arterioles are severely narrowed and in most areas are reduced to a thin white channel (arrow). C. In a CBA/J mouse at P12, there is already extensive loss of the photoreceptors, leaving only a single layer of photoreceptor nuclei (arrow). The inner nuclear and ganglion cell layers are normal. Original magnification × 400. D. In a B6SJL cross, at 3 months of age, the photoreceptors have disappeared and there is focal thinning of the inner nuclear layer (arrow). Original magnification × 400. E and F. Retinal degeneration2. E. At 137 days of age there are large foci of depigmentation (arrow), as well as vascular narrowing. F. The effects of retinal degeneration 2 are much less severe (see C and D) and at 28 months of age, the outer nuclear layer is reduced to one third normal thickness (arrow). However, inner and outer segments remain visible (arrowhead). Original magnification × 400. (A, B, and E, from Hawes, N.L. et al., Mol. Vision, 5:22–29, 1999. With permission.)

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outer segments (reviewed in Chang et al80 and Molday54).22 At 7 days of age, the retinas of homozygous Prph2Rd2 mice are morphologically normal. By P14, the photoreceptor outer segment length lags behind that of normal mice, and by P21 the difference is significant when morphometric measurements are made.85 Narrowing of retinal arterioles occurs as early as P30.83 At 2 months of age by light microscopy, rod outer segments are absent and the outer nuclear layer is reduced to 50% of normal (see Figure 10.4). Progression of the outer retinal degeneration continues with complete loss of both rod and cone nuclei in the peripheral retina by 9 months and in the central retina by 1 year. During this process, the inner retina remains normal, although in older mice there is also cell loss in the inner nuclear layer and intraretinal neovascularization. Patchy loss of pigmentation is seen after three months.83,85 As is true in other retinal degenerations, apoptosis is responsible for loss of the photoreceptors.80 Prph2Rd2/+ mice demonstrate a haploinsufficient phenotype in which the photoreceptor degeneration is more gradual than in homozygotes. Outer segment formation is abnormal, but both inner and outer segments are present at 3 months of age. Photoreceptor nuclei never completely disappear as they do in homozygotes. In addition, the cones are preferentially preserved.55,86

RETINAL DEGENERATION-3 Rd3 differs from Pde6brd1 and Prph2Rd2 mice in that photoreceptor outer segments undergo normal development through P14. However, by P21 loss of both outer segments and photoreceptor nuclei is observed. The degeneration progresses until the outer nuclear layer is totally absent by 10 weeks of age (Figure 10.5). Cone nuclei are the last to disappear.56 Severe narrowing of retinal vessels occurs by P35, although fundus examination is otherwise normal. Involvement of the RPE occurs late with small hypopigmented spots developing about 5 months of age. The pigmentary abnormalities become more prominent by age 10 months.83 The molecular basis for the rd3 mutation is not known.

RETINAL DEGENERATION-4 Rd4 is an autosomal dominant retinal degeneration first described in DBA/2J mice that carry the radiation-induced chromosomal inversion In(4)56Rk. The inversion is a homozygous lethal, but heterozygotes survive and demonstrate an early onset retinal degeneration. There is loss of both outer nuclear and plexiform layers beginning at P10, with complete disappearance by age 6 weeks (see Figure 10.5).57 It appears that the inversion disturbs the Rd4 locus that is located close to the telomeric end of the inversion.87 The gene has not yet been identified.

RETINAL DEGENERATION-5 AND TUBBY-LIKE PROTEIN-1 The rd5 mutant, also known as tub, is characterized by progressive retinal and cochlear degeneration, and by maturity-onset obesity associated with insulin resistance.58,88,89 Tubby mice exhibit many phenotypic similarities to human syndromes. For example, it has been reported that the histopathological changes in the eyes and ears of tubby mice are similar to those seen in individuals with Usher syndrome.58,89 More specifically, tubby appears to be a particularly good model for Usher syndrome Type II because of the similarity in the progressive nature of the disease in both and the lack of vestibular abnormalities.89 The obesity, coupled with the retinal degeneration and hearing loss, also make tubby mice a model for syndromes such as Alstrom and Bardet–Biedl syndromes in humans.90 It has been established that tubby is a loss-of-function mutation.91 Although normal by light microscopy through P14, by P21 there is degeneration of photoreceptor outer segments. This is followed by thinning of the outer nuclear layer that progresses through 8 months of age when all photoreceptors have disappeared. As is so often true, electron microscopy demonstrates that outer segments are never normal. Large membrane-bound vesicles and outer segment fragments are abundant in the subretinal space by 3 to 5 weeks of age (Figure 10.6).58 Cell death occurs by apoptosis.91

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FIGURE 10.5 Retinal degeneration. A to D. Retinal degeneration-3. A. At 150 days of age, fundus photography shows indistinct tiny retinal spots (arrowhead) and some vascular narrowing (arrow). B. By 305 days of age, there are large patches of depigmentation (*) and increased vascular narrowing. C. At 2 months of age, the outer nuclear layer (arrow) and outer segments (OS) are normal. Original magnification × 400. D. In a 26-month-old mouse, the inner and outer segments are absent and only a few photoreceptor nuclei remain (arrow). Original magnification × 400. E and F. Retinal degeneration-4. E. At 3 months of age there are large patches of depigmentation (*) and severe arteriolar narrowing (arrow). F. By 3 months, inner and outer segments have disappeared and there is only a single layer of photoreceptor nuclei (arrow). Original magnification × 400. (A and B, from Hawes, N.L. et al., Mol. Vision, 5:22–29, 1999. With permission.)

The tub gene is a member of a structurally similar family of genes that also includes the tubbylike proteins 1, 2, and 3 (Tulp1, Tulp2, and Tulp3).63-65,92 Null mutants of the Tulp1 gene develop a progressive retinal degeneration similar to tub/tub mice, beginning at P14 with shortening of both inner

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FIGURE 10.6 Retinal degeneration-5. A. Fundus photograph, 3 months of age. Large depigmented areas are present (*) and there is retinal arteriolar narrowing. B. Even by 2 months of age, there is only a single layer of photoreceptor nuclei (arrow). Original magnification × 400. C. A normal wild-type outer retina at 4 weeks of age. The inner (IS) and outer segments (OS) and outer nuclear layer (ONL) are normal. Although the magnification is the same, only about a third of the outer segment length can be seen, an indication of the extent of loss illustrated in D, an affected mouse. D. At 4 weeks of age, both inner (IS) and outer (OS) segments are shortened. The retinal pigment epithelium (PE) is normal. By 9 weeks of age, only a few fragments of outer segments remain (arrow) and the space between the neural retina and pigment epithelium is filled with membranous debris. F. At 3 months there is only an occasional nucleus that might be a photoreceptor (arrows). Otherwise, the inner nuclear layer (INL) lies directly on the pigment epithelium. C to F original magnification × 8100.

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and outer segments of photoreceptors. By 1 month of age, the outer nuclear layer has diminished in thickness and virtually disappears by 20 weeks of age (see Figure 10.6). Electron microscopy demonstrates interphotoreceptor vesicles similar to those seen in tub/tub mice. By 6 months of age, the outer plexiform layer collapses and synaptic complexes have disappeared. Apoptosis is prominent in the outer nuclear layer early in the degeneration.64,65 Mutations within the TULP1 gene have been identified in patients segregating for recessive retinitis pigmentosa.93-95

RETINAL DEGENERATION-6 A characteristic of many human retinal degenerations is the presence of multiple small white retinal spots.11 The autosomal recessive retinal degeneration 6 (rd6) mouse may be a model for the human phenotype known as retinitis punctata albescens, although genetic evidence is lacking at this time. The retina of rd6/rd6 mice develops multiple, small subretinal spots at 8 to 10 weeks of age. The spots are evenly spaced throughout the retina and persist to 22 months of age. Retinal vessels are pale and attenuated by 7 months.60 By light microscopy, the retinas of affected mice appeared relatively normal at P30. However, electron microscopy reveals outer segment breakdown as early as P14 that progresses to loss of inner and outer segments and the outer nuclear layer that is complete by 15 months. The most striking morphological feature of rd6/rd6 mice is the presence of many macrophage-like cells in the subretinal space, between the photoreceptors and RPE. These cells appear as early as 1 week of age and are initially unpigmented with small basophilic nuclei. The cells contain a few pigment granules by P30 and enlarge and become more heavily pigmented with time. It appears that these cells correspond to the white dots seen clinically, as they reach a diameter as great as 35 µm. The cells stain strongly with the monoclonal antibody, MOMA-2 suggesting that they are, indeed, macrophages.60 This is supported by electron microscopy that shows the cells contain numerous phagosomes that surround multiple membrane-enclosed pigment granules and photoreceptor outer segment lamellar debris (Figure 10.7).60

RETINAL DEGENERATION-7 Mice homozygous for the rd7 (Nr2e3rd7) mutation have a pattern of retinal degeneration totally different from that described for retinal degenerations 1 through 6. At weaning age, fundus examination demonstrates many white spots throughout the retina. However, as time progresses, the spots decrease in number while becoming larger and irregular in shape. By 16 months, retinal vessels are attenuated and there is pigment mottling, but the spots disappear.61 Despite the presence of the spots, electroretinograms remain normal until late in the disease after the spots have vanished. The morphological changes correlate well with the clinical findings. The spots are due to the widespread occurrence of prominent folds in the outer nuclear layer that create pseudorosettes reminiscent of those seen in retinal dysplasia. These folds occur as early as P14. One or more lightly pigmented cells that contain cellular debris are usually present in the center of each pseudorosette.61 By 5 months of age, the outer retinal folds begin to disappear, coincident with the decrease in spot number on clinical examination. By 16 months of age, the folds disappear concurrent with a thinning of the outer nuclear layer and outer segments, although neither layer disappears as they do in other retinal degenerations (Figure 10.8).61 Electron microscopy demonstrates that the inner and outer segments of the photoreceptors have become disoriented and “point” toward the center of the pseudorosette. The lightly pigmented cells seen on light microscopy contain pigment granules and cellular debris that suggests phagocytosis. It has not been determined whether these cells are mobilized RPE cells or macrophages, as occurs in rd6 mice. Although the photoreceptors appear by light microscopy to be relatively normal up to 6 months, there is patchy photoreceptor breakdown as early as P30 and this becomes more prominent with increasing age. Similar to the tubby retina, there is extensive formation of interphotoreceptor vesicles in young mice (see Figure 10.8). Rd7 has been identified as a mutation within the Nr2e3 gene.61 Mutations within the same gene has been shown to cause enhanced S-cone syndrome in humans.62 Therefore, Nr2e3 has been postulated to regulate S-cone number.

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PURKINJE CELL DEGENERATION Except for the tubby gene family, the other retinal degenerations described thus far have findings isolated to the eye. However, many human retinal degenerations are part of a broader syndrome that affects other tissues, particularly the central nervous system.11 The Purkinje cell degeneration (pcd) mutant mouse is another example of a syndromic retinal degeneration. Homozygous pcd mice develop

FIGURE 10.7 Retinal degeneration-6. A. Fundus photograph of 2-month-old mouse. Tiny, well-defined white dots are evenly distributed throughout the retina. B. Although the retina appears normal at P21, there is a large macrophage (arrow) in the subretinal space. Original magnification × 630. C. By 8 weeks of age, the outer segments are shortened (compare to B) and fragmented (*). Original magnification × 400. D. By a year of age, a single layer of photoreceptor nuclei (arrow) lies directly on the RPE. Original magnification × 400. E. The inner segments (IS) are normal at 18 weeks of age, but the outer segments (OS) are shortened and fragmented. A large macrophage (M) contains numerous outer segment fragments. F. There is widespread retinal degeneration by 1 year of age. There is intercellular swelling in the outer nuclear layer (ONL). The inner segments (IS) are shortened and swollen and there is also intracellular edema involving the pigment epithelium (PE). E and F original magnification × 6000.

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FIGURE 10.8 Retinal degeneration-7. A. At 3 weeks of age, there are numerous evenly distributed white spots (arrow). B. The white spots in the fundus photograph correspond to numerous folds in the outer retina (arrows). Original magnification × 100. C. The folds are prominent in this 3-month-old mouse and their effects are limited to the outer nuclear layer and photoreceptor elements. Macrophages (arrows) are found within the folds as well as on the surface of the pigment epithelium. Original magnification × 400. D. The folds have nearly disappeared by 4 months of age. The outer segments (arrow) are fragmented and numerous vacuoles are present. Original magnification × 400. E. In a small retinal fold of a 5-week-old mouse, there is marked distortion of the normal position of both inner (IS) and outer (OS) segments. The fold appears centered around a large macrophage (M) in the subretinal space. Original magnification × 8100. F. Folds are no longer present at 16 months of age. The outer segments (OS) are shortened and disorganized. Original magnification × 4000.

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progressively severe retinal degeneration, beginning at P22 followed by the development of ataxia due to loss of cerebellar Purkinje cells between ages 3 and 5 weeks.66 Fundus examination demonstrates a granular appearance by 6 weeks of age and arteriolar attenuation and hypopigmented spots by 10 months.83 This is accompanied by a slow loss of photoreceptors during the first year. Up to P18, photoreceptors appear normal by light microscopy. Beginning at P25, there is disorganization of the outer segments and pyknotic nuclei that are likely due to apoptosis. By P100, while still present, the inner and outer segments are shortened and the outer nuclear layer is only five to six cells thick. By 10.5 months only one to two layers of photoreceptor nuclei remain and by 15.5 months of age nearly all nuclei have disappeared (Figure 10.9). The peripheral retina is more heavily damaged at early stages than the central retina and rods disappear faster than cones. The RPE becomes thinned and depigmented in mice over 1 year age. In addition, invasion of the pigment epithelium by retinal blood vessels occurs. As photoreceptor cells degenerate, their synaptic terminals disappear and the outer plexiform layer becomes thinner. The remainder of the retina is normal, even in older mice. Early in the course of the

FIGURE 10.9 Purkinje cell degeneration. A. At P25 the outer segments (OS) are disorganized and pyknotic photoreceptor nuclei are present. B. By 10.5 months of age, the outer nuclear layer is reduced to one to two rows of photoreceptor nuclei. C. A retinal blood vessel (BV) extends into the RPE. Associated with the vessel is a strand of cells (arrow) displaced from the inner nuclear layer. (From LaVail, M.M. et al., J. Comp. Neurol., 212:217, 1982. With permission of John Wiley & Sons.)

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degeneration, invading cells that are probably macrophages appear in the outer nuclear layer. These cells are initially nonpigmented, but acquire some melanin granules later, presumably from retinal pigment epithelial cells. They also contain abundant lamellar debris from degenerating photoreceptors.66 Electron microscopy of the outer retina of pcd/pcd mice suggests that the initial abnormality is in the photoreceptor inner segment region. At P13, amorphous cytoplasmic densities are found in Muller cells and a small number of interphotoreceptor vesicles are found adjacent to the inner segments. Swelling of Muller cells and photoreceptor inner segments became increasingly prominent from P13 to P25. The number of vesicular profiles also increase during this time. Rod outer segments and RPE remain normal through P25, but begin to degenerate thereafter. By 5.5 months only a few disorganized photoreceptor outer segments can be identified. Inner segments can still be identified and there is a persistence of the vesicular profiles. By 15 months of age, all photoreceptor nuclei, inner, and outer segments have disappeared.67

NERVOUS Nervous mutant mice (nr/nr) exhibit juvenile hyperactivity and ataxia and lose 90% of cerebellar Purkinje cells during the first 7 weeks of life. Retinal degeneration proceeds at a much faster rate than in pcd mice. Fundus examination demonstrates large white spots that become confluent by 7 months of age (Figure 10.10).83 As early as P11, increased numbers of pyknotic nuclei are seen, presumably due to apoptosis. Outer segments never reach normal length and quickly become disorganized. During the same time period, the thickness of the outer nuclear layer decreases to 60% that of normal mice. After this acute phase, the rate of degeneration slows, but by 7.5 months, there is only a single layer of photoreceptor nuclei. In older mice, the outer nuclear layer disappears in places and inner retinal vessels invade the outer retina and RPE, as happens in pcd mice.68,96 Electron microscopy (Figure 10.11) can detect changes in nr/nr mice as early as P6. Mitochondria of the photoreceptor inner segments are two to three times normal size. By P9, affected mice lag behind normal mice in outer segment differentiation. At the same time, affected mice show a decrease in free ribosomes, rough-surfaced endoplasmic reticulum, and Golgi. At P16, the outer segments are shorter than normal and often disorganized. By 5 months small amounts of lamellar membranous material separate the inner segments from the pigment epithelium. Ultimately, photoreceptor nuclei lie in direct contact with the pigment epithelium. The inner retina and RPE remain normal, although there are an increased number of phagosomes identified during the first few months of life.69

NEURONAL CEROID LIPOFUSCINOSES Many different neuronal ceroid lipofuscinoses (NCL) occur in humans. Most have an onset in infancy, although some are first identified in adults. The most common phenotypes include progressive dementia, seizures, and retinal degeneration (reviewed in Spencer11 and Cooper et al.97). Two mouse models with clinical and morphological phenotypes similar to human NCL have been reported. The motor neuron degeneration mutant mouse (mnd) develops paresis of the hind limbs beginning at 5 months of age and progresses to paralysis by 14 months. This is due to accumulation of membranous intracellular lipid deposits in multiple central nervous system locations.70,71,97 As is true in human disease, mnd/mnd mice develop progressive retinal degeneration. Fundus examination demonstrates early arteriolar attenuation by 6 weeks of age and a granular appearance of the RPE. Photoreceptor development in mnd mice is normal until P35 when the thickness of the outer nuclear layer begins to decrease and completely disappears by 6 months of age. The peripheral retina is affected earlier than the posterior retina. In addition to the loss of photoreceptors, all cells of the retina and pigment epithelium contain membrane-bound, lysosome-like inclusions with curvilinear profiles similar to those seen in NCL.70,71

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Similar findings occur in the neuronal ceroid lipofuscinosis mouse (nclf) that maps to mouse Chr 9 in an area homologous with human 15q21 where late infantile NCL (CLN6) has been mapped (reviewed in Bronson et al.72). Progressive paralysis of somatic muscles is accompanied by retinal degeneration. White retinal spots develop around the fourth month of life. There is loss of the outer nuclear layer and photoreceptor outer segments that progress slightly more slowly than in mnd mice. Lamellar cytoplasmic inclusions are abundant in the eye and central nervous system.72

FIGURE 10.10 Retinal degeneration in nervous mutant mice. A. Wild-type mouse at P16. The outer nuclear layer and inner and outer segments are normal. B. In a P16 nr/nr mouse, the outer nuclear layer is much thinner than in the wild-type mouse and contains numerous pyknotic nuclei (arrows). Both pictures taken at the same magnification. C. Retina at age 2.5 months. A few normal-diameter outer segments reach the surface of the RPE (arrowheads), but most are disorganized and arranged in membranous whorls (W). D. Retina at 7.5 months of age. The outer nuclear layer (arrow) is reduced to a single row of nuclei that lie directly on the apex of the RPE cells. Inner and outer segments are absent. E. The RPE is invaded by retinal capillaries (C). (From LaVail, M.M. et al., J. Comp. Neurol., 333:168, 1993. With permission of John Wiley & Sons.)

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FIGURE 10.11 Retinal degeneration in nervous mutant mice. A. In a wild-type mouse at P16, the outer segments (OS) are regularly aligned. The inner segments (IS) contain abundant membranes of the Golgi complex (G) and rough endoplasmic reticulum (rer), as well as many polyribosomes (p). The mitochondria are small and elongated. B. In a P16 nr/nr mouse, the outer segments are shorter than normal and in some the membranes are arranged in disorganized arrays. Some outer segment tips are apposed to the apical surface of the RPE in relatively normal configuration (arrows), while some membranous whorls are also found (arrowhead). There is very little Golgi or endoplasmic reticulum and mitochondria are shorter and enlarged in diameter. C. At age 5.5 months, large lamellar whorls (W) are present in the outer segment zone. These whorls, along with disorganized outer segment membranes and inner segments (IS), are surrounded by apical processes of the RPE (arrowheads) that extend as much as halfway through the photoreceptor layer. Microvillous processes of Muller cells (mv) extend into the photoreceptor layer. The external limiting membrane zonulae adherentes (za) are intact. D. At 7.5 months of age, the outerment membranes have disappeared and the inner segments (IS) are reduced in length and contain large rounded mitochondria (m). (From White, M.P. et al., J. Comp. Neurol., 333:182, 1993. With permission of John Wiley & Sons.)

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CONE-ROD HOMEOBOX GENE A number of human ocular diseases have been associated with the cone-rod homeobox gene (Crx), including Leber congenital amaurosis,98 autosomal dominant cone rod dystrophy,99,100 and retinitis pigmentosa.101 In the eye, Crx is expressed only in developing and mature photoreceptor cells. Crx regulates photoreceptor differentiation,73 as demonstrated by the failure of Crx-deficient mice to develop photoreceptor outer segments. By P21, there is a reduction in thickness of the outer nuclear layer and by 6 months only one to two layers of photoreceptor nuclei remain. The inner retina and retinal pigment epithelium are unaffected (Figures 10.12 and 10.13).74

FIGURE 10.12 CRX-deficient mice. A (P14) and D (P21) are wild-type mice. The outer nuclear layer, inner segments (IS), outer segments (OS), and retinal pigment epithelium (PE) are all normal. B (P14) and E (P21) are Crx+/– (heterozygous) mice. The outer segments are clearly shorter than in wild-type mice at P14, but are similar in length by P21. C (P14) and F (P21) are Crx–/– mice. There are no outer segments at either age and by P21 there are decreased numbers of photoreceptor nuclei. (From Cepko, C., Nat. Genet., 23:466–70, 1999. With permission.)

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FIGURE 10.13 Phenotypes of CRX-deficient mice at 1 to 6 months. A. Dark-adapted rod and light-adapted cone ERG responses from 4-week-old wild-type, Crx+/–, and Crx–/– mice. Response amplitudes (µV) of b-waves are indicated below waves with standard error. No significant amplitude was detected in ERGs from homozygous mice. B. ERGs from 2-month-old mice. C. ERGs from 6-month-old mice. D. At 6 months of age, wildtype mice have normal retinal morphology. E. At 6 months of age, Crx+/– mice show early photoreceptor degeneration. F. At age 2 months in Crx–/– mice, there is loss of inner and outer segments, but the remainder of the retina is normal. G. At 6 months of age, the photoreceptor layer consists of only two to four rows of cells in Crx–/– mice. (From Cepko, C., Nat. Genet., 23:466–70, 1999. With permission.)

ARRESTIN—OGUCHI’S DISEASE Rhodopsin kinase and arrestin have both been implicated in stationary night blindness (Oguchi’s disease) in human eyes.102 Since human anatomic material is scarce, information regarding morphological alterations for this disease is limited.11 An arrestin knockout model of Oguchi’s disease in mice demonstrates enhanced susceptibility to light exposure. When arrestin-null mice are exposed to cyclic light, there is a loss of photoreceptors beginning at 100 days of age. By 1 year, fewer than 50% of photoreceptors survive. The changes are more severe and accelerated when these mice are exposed to constant light.75

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RHODOPSIN MUTATIONS The rhodopsin gene in humans is highly mutable and over 70 mutations have been described that are associated with retinitis pigmentosa.40 Transgenic mice that carry some of these mutations have been generated in an effort to understand the mechanisms through which a mutant rhodopsin gene leads to retinitis pigmentosa. Mice that carry a human rhodopsin mutation, pro23his, develop severe retinal degeneration by P20 with loss of inner and outer segments as well as photoreceptor nuclei. The inner retina is normal, but the RPE demonstrates variation in cell size and pigmentation corresponding to the fundus appearance.103 Similar morphologic findings characterize both thr17meth and pro347ser rhodopsin mutations in mice.104,105

OTHER RETINAL DEGENERATIONS The goal of this section on retinal degenerations has been to provide information on the morphological patterns associated with these diseases as well as to describe the more common, well-characterized mutations. Targeted mutations directed toward a specific retinal cell type can produce many variations on what has already been described. Cones,106 rods,107 and horizontal cells108,109 have all been targeted by specific transgenes and induced to degenerate, often with consequences for the surrounding cells. These, as well as other examples, will not be discussed because the morphological changes are similar to those degenerations already reviewed.

RETINAL DETACHMENT Although a common problem in human eyes,11 retinal detachment is unusual in mice. Because formalin-fixed and paraffin-embedded eyes are particularly prone to artifactual retinal detachment, this artifact must be properly identified. Because there is a potential space between the retina and RPE, these layers are easily separated. For this reason, opening the unfixed eye to promote fixation is best avoided. The same caution applies to fixation for electron microscopy (see Chapter 13). There are two features useful in distinguishing true detachment from an artifact: photoreceptor degeneration and subretinal fluid. The high metabolic rate of the photoreceptors and their total dependence on the choriocapillaris causes outer and inner segments to degenerate within several hours after separation of the retina from the pigment epithelium. Proteinaceous fluid frequently accumulates beneath the retina and is strongly eosinophilic. When the detachment has persisted for a few weeks, macrophages or mobilized retinal pigment epithelial cells accumulate in the subretinal space and in the subretinal fluid. The retina may also be detached by subretinal hemorrhage or there may be bleeding into a previously existing detachment (Figure 10.14). Secondary retinal detachment is frequently a feature of vitreous traction due to vitreous fibrosis after vitreous hemorrhage, persistent hyperplastic primary vitreous, and those mutations in which the hyaloid vessels persist or proliferate, causing a traction retinal detachment (see Chapter 9). A specific example of vitreous traction occurs in mice that express platelet-derived growth factor (PDGF). There are two forms of PDGF, PDGF-A and PDGF-B. When the rhodopsin promoter is coupled to either form, both are expressed in the retina, although the phenotypic results are quite different. Mice transgenic for PDGF-A develop intraretinal gliosis that appears to protect them from oxygen-induced ischemic retinopathy.110 In contrast, when PDGF-B is overexpressed, proliferation of epiretinal and subretinal membranes occurs and the retina often became attached to the posterior lens surface. In the detached retina of these mice, there is diffuse degeneration of all retinal layers, intraretinal cyst formation, and migration of RPE cells into the retina. The production of intraretinal astrocytes, pericytes, and vascular endothelial cells is also evident. The traction in these mice is the result of contraction of epiretinal and subretinal membranes.

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FIGURE 10.14 Retinal detachment and drusen. A. Retinal detachment of unknown etiology in an older mouse. The optic nerve is barely visible (arrow). There is a bullous retinal detachment (*). The retina is white, because the normal pattern of the retinal pigment and epithelium and choroid are obscured. B. The presence of separation of the retina from the pigment epithelium as seen here does not always mean a retinal detachment is present. This picture shows an artifact of preparation. Note that there is no subretinal fluid and that the inner segments (arrow) and outer segments (arrowhead) are well preserved (compare to C). Original magnification × 400. C. This represents a long-standing retinal detachment, similar to that shown in A. Although there is some suggestion of inner segments (arrow), the outer segments are absent. A large amount of proteinaceous subretinal fluid (SRF) is present, and there are numerous phagocytic cells (arrowhead) in the subretinal space. Original magnification × 400. D. One mechanism of retinal detachment is vitreous traction. In a p53-null mouse, the fibrovascular persistent hyperplastic primary vitreous (arrow) has pulled the retina up to the posterior lens capsule. Original magnification × 100. E. Retinal drusen are frequently found in human eyes, but are rare in mice. Here, the retina has degenerated over the apex of a large drusen (arrow). Original magnification × 400. F. At higher power, the lamellar pattern of the drusen (arrow) is apparent. The RPE is disrupted and the outer retina has degenerated. Original magnification × 630.

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PRIMARY DISEASES OF THE RETINAL PIGMENT EPITHELIUM As has already been implied, separation of diseases of the retina from those of the pigment epithelium is difficult because of their anatomic, embryological, and physiological interrelationship.111,112 The effects of PDGF, described in the previous section, are one example.113 It is established that many specific factors expressed during development have reciprocal functions between these two cell layers (for review, see Jean et al.114), including fibroblast growth factor (FGF) and its receptors,2,115 microphthalmia-associated transcription factor,24 bone morphogenetic proteins,116 and members of the Pax gene family.117 In the preceding section, many of the primary retinal degenerations also involve the pigment epithelium. This section will be limited to examples in which the pigment epithelium is the initial tissue affected.

DOMINANT NEGATIVE FGF RECEPTOR-1 When targeted to the RPE, there is strong expression of a dominant negative FGFR-1 in developing mouse RPE. Homozygotes develop severe microphthalmia with maldevelopment of many ocular structures (Figure 10.15). Hemizygotes have decreased numbers of choroidal melanocytes and capillaries. The RPE is flattened and Bruch’s membrane is absent. Contact of pigment epithelial microvilli with outer segments is reduced compared to wild-type mice. The outer segments are shorter than normal and, in older mice, the outer nuclear layer is only 50% of normal thickness. The decrease in the choriocapillary bed raises the question of the role of circulatory insufficiency in this model.115

GYRATE ATROPHY In humans, gyrate atrophy is an uncommon autosomal recessive chorioretinal degeneration due to a deficiency of ornithine δ-aminotransferase (OAT). It is characterized by large patches of well-demarcated circular fundus lesions with hyperpigmented edges and with hyperornithinemia (reviewed in Wang et al.118). OAT is strongly expressed in both the neural retina and RPE.119 OAT-deficient mice develop normally for the first 60 days of life, but then develop sporadic necrosis of RPE cells that show pale cytoplasm and swollen mitochondria. By 6 months of age this progresses to diffuse pigment epithelial involvement with many large phagosomes as well as lipidlike inclusion bodies. Concurrently, there is severe fragmentation of the photoreceptor outer segments. Photoreceptor nuclei also gradually decreased, most likely by apoptosis.118,120 The retinal degeneration in OAT-deficient mice can be completely eliminated by maintaining the animals on an arginine-restricted diet, which relieves the hyperornithinemia.121

SENESCENCE-ACCELERATED MOUSE Although age-related macular degeneration is a major visual problem in humans,122,123 there are few examples of these problems in mice. The multiple strains of senescence accelerated mice (SAM) (reviewed in Majji et al.124) are an important exception. Different substrains of these mice develop different ocular diseases including corneal scarring, cataract, persistent hyaloid vessels, and periocular lymphomas.124-127 The SAM P8 strain also develops severe changes in the retinal pigment epithelium and Bruch’s membrane after 10 months of age. Bruch’s membrane becomes diffusely thickened and, in addition, develops focal thickening adjacent to the base of the RPE. Scattered pigment epithelial cells are depigmented. Electron microscopy demonstrates disruption of pigment epithelial microvilli as early as 8 months of age. The cytoplasm of pigment epithelial cells becomes filled with electronlucent vacuoles that may possibly be lipid in nature. Bruch’s membrane is three to four times normal thickness and demonstrates localized deposits of basal laminar material. There is minimal atrophy of the choriocapillaris. Despite RPE and Bruch’s membrane changes that are reminiscent of human agerelated macular degeneration, the neural retina is normal.124

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FIGURE 10.15 Transgenic mice with dominant-negative FGF receptor. A and D. In adult wild-type mice, retina, choroid, RPE, and Bruch’s membrane (arrow) are well developed. Melanosomes are abundant. B and C. In transgenic mice, there is marked thinning of the choroid with a reduction in melanocyte number. E. In a transgenic mouse, Bruch’s membrane is absent (arrow) and RPE microvilli fail to make contact with photoreceptor outer segments (*). There are decreased numbers of choroidal melanocytes. F. Bruch’s membrane is hypoplastic (arrow), the basal infoldings of the RPE (arrowhead) are disorganized. G. In a severely affected mouse, photoreceptor inner and outer segments are absent, the pigment epithelium is flattened, Bruch’s membrane and the choriocapillaris are missing, and there are no melanocytes in the choroid. (From Javerzat, S., Exp. Eye Res., 71:395–404, 2000. With permission of Harcourt.)

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DEGENERATIVE AND REACTIVE CHANGES IN THE RPE Coincidental changes in the degree of pigmentation of the pigment epithelium occur in many retinal degenerations. Primary changes in pigmentation are important in the platelet storage pool diseases and in induced tumors of the RPE, which were discussed in Chapter 9. The RPE may also be altered in colobomas that affect the choroid, retina, and optic nerve (see Chapter 11). Focal disturbance of

FIGURE 10.16 Retinal pigment epithelium. A. A chorioretinal adhesion of undetermined etiology (between arrows) binds the two layers together. Original magnification × 200. B. At higher magnification, the fibrovascular nature (*) is more apparent. The adhesion interrupts the RPE (arrowhead). Original magnification × 400. C. The RPE has proliferated (*), separating the retina from the choroid and disrupting the normal structure of the pigment epithelium. Original magnification × 400. D. RPE proliferations may reach large size and when heavily pigmented (arrow) could be mistaken for a benign or malignant tumor. Original magnification × 400. E. True hyperplasia of the RPE is usually a developmental abnormality. Note the junction (arrow) between hyperplastic RPE on the left and normal RPE on the right. Original magnification × 400. F. At higher magnification, the small area of normal RPE (between arrows) is less than half the height of the hyperplastic RPE. Original magnification × 630.

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the RPE may be due to trauma or inflammation. The end result is usually a focal chorioretinal scar that binds the two tissues together (Figure 10.16). Although uncommon, the RPE may undergo a reactive form of proliferation following injury or chronic retinal detachment. Because it is a relatively undifferentiated group of cells, the RPE retains the ability to produce fibrous tissues as well. In some instances, the fibrous tissue can be part of a multilayer benign proliferation of RPE cells that forms a pseudotumor (see Figure 10.16). Congenital hyperplasia of the RPE is often observed in human patients11 and represents a true hyperplasia including both increased cell size and increased melanosome size. This has not been reported in mice as a developmental lesion, but a phenocopy has been observed in the retinas of older mice that are homozygous for Pde6brd1 (see Figure 10.16).

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106. Ying, S. et al., Retinal degeneration in cone photoreceptor cell-ablated transgenic mice, Mol. Vis., 6:101, 2000. 107. Peachey, N.S. et al., Functional consequences of oncogene-induced photoreceptor degeneration in transgenic mice, Vis. Neurosci., 12:513, 1995. 108. Hammang, J.P. et al., Oncogene expression in retinal horizontal cells of transgenic mice results in a cascade of neurodegeneration, Neuron, 10:1197, 1993. 109. Peachey, N.S. et al., Functional consequences of oncogene-induced horizontal cell degeneration in the retina of transgenic mice, Vis. Neurosci., 14:627, 1997. 110. Yamada, H. et al., Platelet-derived growth factor-A-induced retinal gliosis protects against ischemic retinopathy, Am. J. Pathol., 156:477, 2000. 111. Raymond, S.M. and Jackson, I.J., The retinal pigmented epithelium is required for development and maintenance of the mouse neural retina, Curr. Biol., 5:1286, 1995. 112. Morrow, E.M., Furukawa, T., and Cepko, C.L., Vertebrate photoreceptor cell development and disease, Trends Cell Biol., 8:353, 1998. 113. Campochiaro, P. et al., Platelet-derived growth factor is an autocrine growth stimulator in retinal pigmented epithelial cells, J. Cell Sci., 107:2459, 1994. 114. Jean, D., Ewan, K., and Gruss, P., Molecular regulators involved in vertebrate eye development, Mech. Dev., 76:3, 1998. 115. Rousseau, B. et al., Neural and angiogenic defects in eyes of transgenic mice expressing a dominantnegative FGF receptor in the pigmented cells, Exp. Eye Res., 71:395, 2000. 116. Mathura, J.R. et al., Bone morphogenetic proteins-2 and -4: negative growth regulators in adult retinal pigmented epithelium, Invest. Ophthalmol. Vis. Sci., 41:592, 2000. 117. Schwarz, M. et al., Spatial specification of mammalian eye territories by reciprocal transcriptional repression of Pax2 and Pax6, Development, 127:4325, 2000. 118. Wang, T. et al., A mouse model of gyrate atrophy of the choroid and retina, J. Clin. Invest., 97:2753, 1996. 119. Hotta, Y. and Kato, T., Ornithine aminotransferase distribution in ocular tissues and retinas of cat and mouse, Invest. Ophthalmol. Vis. Sci., 30:1173, 1989. 120. Wang, T. et al., Mice lacking ornithine-δ-amino-transferase have paradoxical neonatal hypoornithinaemia and retinal degeneration, Nat. Genet., 11:185, 1995. 121. Wang, T. et al., Correction of ornithine accumulation prevents retinal degeneration in a mouse model of gyrate atrophy of the choroid and retina, Proc. Natl. Acad. Sci. U.S.A., 97:1224, 2000. 122. Klein, R. et al., The Wisconsin epidemiologic study of diabetic retinopathy, Arch. Ophthalmol., 112:1217, 1994. 123. Retinal Diseases Panel, Vision Research, A National Plan: 1999–2003, U.S. Department of Health and Human Services, Washington, D.C., 1999. 124. Majji, A.B. et al., Age-related retinal pigment epithelium and Bruch’s membrane degeneration in senescence-accelerated mouse, Invest. Ophthalmol. Vis. Sci., 41:3936, 2000. 125. Hosokawa, M. et al., Persistent hyaloid vascular system in age-related cataract in a SAM strain of mouse, Exp. Eye Res., 57:427, 1993. 126. Hosokawa, M. and Ueno, M., Aging of blood–retinal barrier and neuronal cells of eye and ear in SAM mice, Neurobiol. Aging, 20:117, 1999. 127. Shoji, M. et al., Senile ocular amyloidosis in SAM and BALB/c strains of mice, Mech. Ageing Dev., 120:87, 2000.

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Optic Nerve and Orbit Richard S. Smith, Simon W. M. John, and John P. Sundberg

CONTENTS Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .227 Optic Nerve—Developmental Abnormalities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .228 Cyclopia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .228 Optic Nerve Aplasia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .228 Optic Nerve Hypoplasia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .229 Optic Nerve Coloboma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .229 Dysmyelination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .231 Demyelination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .234 Copper Deficiency—Menkes Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .234 Optic Nerve Degenerations and Inflammation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .234 Wallerian Degeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .234 Axonal Degeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .235 Optic Neuritis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .235 Orbit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .235 Developmental Abnormalities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .235 Lacrimal Gland—Sjogren’s Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .235 Thyroid Orbitopathy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .237 Orbital Inflammation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .237 Neoplasms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .239 Neurofibromatosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .239 Lymphoproliferative Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .229 Lacrimal and Harderian . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .240 Myoepithelioma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .241 Fibro-Osseous Proliferation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .241 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .244

INTRODUCTION Ocular diseases such as glaucoma and retinal degeneration exert indirect effects on the optic nerve, and these specific problems are discussed in other chapters. One purpose of this chapter is to review those conditions in which the optic nerve is the primary target. The optic nerve is surrounded within the orbit by a variety of tissues, including the Harderian glands, smooth and striated muscle, cartilage, fat, and connective tissue. Despite the complex array of orbital tissues, relatively few abnormalities have been described in mice; these will be discussed at the end of this chapter.

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OPTIC NERVE—DEVELOPMENTAL ABNORMALITIES CYCLOPIA Under normal circumstances, the orbit and midline facial structures are recognizable early in embryonic development. Fusion of the optic vesicles to form a single midline eye results in cyclopia, a malformation associated with central nervous system, spinal, and skeletal abnormalities generally incompatible with life. Cyclopia has been produced in mice by targeted disruption of the sonic hedgehog (Shh) gene. Shh–/– mice die around the time of birth and demonstrate failure of development of the facial structures, multiple skeletal and central nervous system defects, and cyclopia (Figure 11.1). Both cyclopia and holoprosencephaly occur after exposure to the Veratrum alkaloid cyclopamine that acts specifically to inhibit sonic hedgehog signal transduction.1,2 The internal ocular structures do not develop because the optic vesicle never invaginates. The optic nerve is absent. Absence of Shh disrupts expression of many developmental genes, which undoubtedly contributes to the multiple abnormalities characteristic of these mice.3

OPTIC NERVE APLASIA The earliest reports of optic nerve aplasia came from a strain originally found and interbred by Dr. C. C. Little in 1938 at The Jackson Laboratory. Strain ZRDCT-An mice have no eyes 80 to 90% of the time, and the remaining 10 to 20% exhibit microphthalmia.4 Although not characterized, the eye findings are due to a single recessive gene.5 In addition to the smaller eye, the microphthalmic

FIGURE 11.1 Cyclopia. A and B. E9.5 embryos hybridized with microphthalmia (Mitf) antisense RNA. A. Wild-type embryo sectioned at level of optic vesicles (white arrows). The forebrain is indicated by a white asterisk. B. In a Shh–/– embryo, only a single optic vesicle is present. (From Beachy, P.A., Nature, 383:407–413, 1996. With permission.)

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subgroup has aplasia of the optic nerve.6 From a morphological viewpoint, optic nerve aplasia is always related to aberrant formation or faulty closure of the optic (choroidal) fissure. When the optic vesicle invaginates, the optic fissure forms ventrally around E117 and provides the pathway for the hyaloid artery to enter the eye. Nerve fibers from the developing retina enter the optic fissure and grow back along its walls to the brain, and by E13, the fissure closes (reviewed in Silver and Robb8). Closure of the fissure is associated with prominent apoptosis along the developing area of fusion.9 In ZRDCT-An mice with microphthalmia the optic fissure does not form completely and often remains open. As a result, normal developing nerve fibers from retinal ganglion cells fail to exit from the eye. The failure of nerve fiber migration leads to a nearly total loss of retinal ganglion cells in older mice. The remainder of the retina is normal except for occasional foci of retinal dysplasia.6 Optic nerve aplasia also occurs in mice homozygous for the ocular retardation gene (originally or, now Chx10or). The eyes develop normally until E11 but are slightly smaller than normal littermates. By E14, in addition to the smaller eye, the retina is thin and the vitreous cavity and lens are small for the age of the mice. The optic fissure closes completely, blocking the developing nerve fibers from leaving the eye. Accordingly, the optic chiasm is absent. The total occlusion of the optic fissure prevents development of the ocular circulatory system.8,10 Another allele, Chx10or-J, is a null mutation of the Chx10 homeobox gene.11,12

OPTIC NERVE HYPOPLASIA Development of the optic nerve depends not only on the proper sequence of events during formation and closure of the optic fissure, but also on a complex set of proteins responsible for directing the migration of developing nerve fibers from the retina.13,14 During development (reviewed in Deiner et al.13), axons arising from the retinal ganglion cells are directed toward the optic nerve, enter it, and extend back to make connections in the brain (Figure 11.2). The protein netrin-1 and its receptor DCC (deleted in colorectal cancer) play important roles in early axon pathfinding. Mice deficient in netrin-1 or DCC have hypoplastic optic nerves. This is apparently due to a failure of axons to enter the optic nerve after they reach the optic disc.13 Optic nerve hypoplasia also occurs as a secondary effect of prenatal exposure to alcohol. When intraperitoneal alcohol is administered to pregnant mice around day 7 of embryonic development, there is a high incidence of microphthalmia15,16 and optic nerve hypoplasia.17-19 The reported findings are similar to what has been observed in human infants exposed to maternal alcohol during pregnancy.20,21 The intraocular effects of fetal alcohol syndrome in mice were discussed in Chapter 8. The optic nerve hypoplasia model uses offspring that were exposed to alcohol at E12. These mice have a delay in myelination compared to controls.19 Shortly after birth, the size of the optic nerve is comparable between treated and untreated mice. Between 2 and 4 months of age axon numbers decrease in cross-sectional area in optic nerves of treated mice. It has been suggested that this loss is due to changes in neurotrophic factors, but it may also be explained as delayed degeneration of previously injured axons.17,18

OPTIC NERVE COLOBOMA Colobomas of the optic nerve are developmental defects in which a portion of the optic nerve is absent. Colobomas may be partial or complete and are frequently accompanied by colobomas of the retina and choroid. They are an uncommon defect in human eyes and in some cases may be accompanied by late subretinal neovascularization.22-28 Colobomas of the optic nerve and retina in mice (see also Chapter 8) are found in many mutations, as summarized in Table 11.1. This list suggests the complexity of molecular mechanisms responsible for tissue remodeling and axonal migration that interact during formation of the optic nerve. These complex interacting

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FIGURE 11.2 Optic nerve hypoplasia and colobomas. A. In a mouse with a large coloboma of the optic nerve (similar to E), the “optic nerve” (ON) is missing the typical arrangement of nerve fibers that normally run parallel to the nerve surface. Original magnification × 200. B. A thin dural sheath of the nerve is present (arrow), but there are no nerve fibers, which should be seen easily at this magnification. The nuclei are probably those of astrocytes and oligodendrocytes. Original magnification × 400. C. A coloboma of the optic nerve and retina (between the arrows) is much larger than the normal size of an optic nerve (between arrowheads). (From Hawes, N.C. et al., Mol. Vision, 5:22–29, 1999. With permission.) D. In another large coloboma, there is no recognizable optic nerve tissue in the scleral foramen (arrows). Retinal tissue (R) has extended into the space normally occupied by the optic nerve. The typically thick, well-defined peripapillary nerve fiber layer is absent (arrowheads). Original magnification × 200. E. The retina adjacent to the optic nerve is thin and retinal layers are missing (arrows) in a large coloboma of nerve and retina. Original magnification × 100. F. In a higher magnification, the normal substance of the optic nerve is replaced by fibroglial tissue (*). Original magnification × 400.

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Table 11.1 Mouse Mutations with Coloboma Name

Chromosome

Ref.

Mitf

6

29

Bst

16

30

Pax21Neu

19

31

Coloboma

Cm

2

32

Total cataract and microphthalmia

Tcm

4

33

—

2, 11, 14, 15

34–36

Gli3Xt

13

37, 38

Microphthalmia Belly spot and tail Neuherberg 1 allele of paired box gene 2

Retinoic acid receptor mutations Extra toes

Symbol mi

molecular mechanisms are beyond the scope of this book. The interested reader is referred to additional basic and review articles as an introduction to these developmental issues.14,39-42 Morphologically, colobomas have a similar appearance regardless of the specific genetic abnormality. In addition to effects on the optic nerve, the adjacent retina, choroid, and retinal pigment epithelium (RPE) are often morphologically altered by a coloboma. In minimally affected mice, decreased numbers of nerve fibers enter the optic nerve and the retina and choroid may be slightly truncated in the peripapillary region. With more severe involvement, the normal structure of the optic nerve is lost, but may still be recognizable. Nerve fibers are usually absent. Choroidal melanocytes, RPE, and retina may prolapse into the empty optic nerve (see Figure 11.1). This tendency is more pronounced in the most severe cases in which the optic nerve is completely absent. While uncommon, there may also be colobomas that involve retina or choroid. Focal retinal dysplasia and areas of incomplete retinal differentiation are frequent in colobomas. Age-related subretinal neovascularization occurring in Bst/+ mice with colobomas is discussed in Chapter 10.30

DYSMYELINATION Myelin acts to insulate axons of the central and peripheral nervous systems. Myelin in the central nervous system consists of myelin basic protein (gene-Mbp) and proteolipid protein (gene-Plp) and its isoform, DM20. Mutations with the colorful names jimpy and rumpshaker represent two of several alleles of the X-linked Plp gene.43-45 Mutations in the Plp gene are responsible for Pelizaeus–Merzbacher disease (PMD) and X-linked spastic paraplegia type 2 in humans.46 Mouse Plp mutations are models of these diseases. In humans, PMD is associated with demyelination and atrophy of the optic nerve.47,48 Plp-deficient mice fail to produce myelin of normal thickness and morphology. Between 6 and 8 weeks of age focal axonal swellings appear that contain dense bodies, multivesicular bodies, and mitochondria. As these swellings enlarge, the abnormal myelin sheath becomes thinner and eventually disappears, axons degenerate, and oligodendrocytes disappear.46,49 Electron microscopic evaluation demonstrates that the oligodendrocytes never properly differentiate and undergo programmed cell death (Figures 11.3 and 11.4).50 The molecular mechanisms remain unclear, but may relate to toxic effects arising from misfolded PLP protein.51

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FIGURE 11.3 PLP-DM20 deficiency. Axonal spheroids containing membranous organelles. A. Transverse section of spinal cord. B. Longitudinal section of optic nerve of Plp-/Y mouse. A and B. Axonal swellings were often in the paranodal region adjacent to the node of Ranvier (N). Large spheroids in the optic nerve of a 12month-old Plp-/Y mouse. C. The swellings are primarily neurofilamentous while those in D contain mainly membranous organelles. The myelin sheath is in the process of degeneration. E. Mice deficient for both MBP and PLP show spheroids identical to those of PLP-deficient mice, although they are absent in MBP-deficient (shiverer) mice. (Illustrations courtesy of Klaus-Armin Nave.) (From Griffiths, I. et al., Science, 280:1610, 1998. With permission of the American Association for the Advancement of Science.)

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FIGURE 11.4 A. Cerebellar cortex from Plp-/Y mouse immunostained for phosphorylated neurofilaments shows swollen Purkinje cell axons (arrows). The cell body of one cell (open arrow) contains phosphoylated neurofilaments suggesting that its axon has degenerated. B. Degenerating (arrow) or swollen (open arrow) smallcaliber fibers in the fasciculus gracilis in a 22-month-old Plp-/Y mouse. The density of axons is decreased, compared to C, which is from the spinal cord of the same mouse. D. Proximal optic nerve of a 22-month-old Plp-/Y mouse, showing the optic disc (O), the lamina cribrosa (to left of large arrow), and the myelinated portion of the nerve (to right of small arrow). The small arrow points to an axonal swelling. E. Normal unmyelinated axon from lamina cribrosa area. F. A myelinated optic nerve axon shows abnormal organelle accumulation. (Illustrations courtesy of Klaus-Armin Nave.) (From Griffiths, I. et al., Science, 280:1610, 1998. With permission of the American Association for the Advancement of Science.)

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DEMYELINATION Since PLP is a major myelin lipid, deficient mice are presumed never to develop normal myelin structures and this is appropriately termed dysmyelination. In contrast, if myelin is normally formed and then lost in adult life, the process is one of demyelination. When thinking of animal models, this is an important distinction, since demyelinating diseases such as multiple sclerosis in humans are a major health problem. A mouse model of demyelination has been produced by transgenic insertion of 70 copies of the gene that encodes DM20, the isoform of PLP. DM20 has an important role in myelogenesis, but is expressed less once maturity is reached. In these transgenic mice, DM20 expression persists into adult life.46,51,52 These mice are clinically normal through 3 months of age and then develop progressively severe tremors and unsteady gait with death by 8 to 10 months of age. At death, affected mice have less than 20% of the normal amount of myelin and demonstrate prominent astrogliosis, the latter characteristic of demyelinating diseases. By electron microscopy, myelin is disrupted and myelin debris is located in astrocytes. This is accompanied by a mild lymphocytic infiltrate. The optic nerves are also affected.52 There is actually decreased myelin synthesis in the transgenic mice with multiple copies of DM20 mice that demonstrate increased amounts of DM20, increased fatty acylation, and decreased amounts of proteolipid protein.53 While these mice undergo demyelination, it appears that myelogenesis is disturbed at younger ages, so ND4 mice may belong in the dysmyelination category. This conclusion is supported by the dysmyelination changes observed in PLP/DM20 transgenic mice that carry extra copies of the gene.54

COPPER DEFICIENCY—MENKES DISEASE Menkes disease is an X-linked human disease characterized by epilepsy, kinky hair, hypopigmentation, and optic atrophy. The metabolic basis of Menkes disease is defective copper transport due to mutations in the Cu2+-adenosinetriphosphatase gene (Atp7).45,55,56 Studies of humans and mice demonstrate many different alleles of the Atp7a gene that include deletions, insertions, and point mutations.45,57 In mice, the mottled allele (Atp7aMo) is the mutant homologue of human ATP7A. The mouse macular allele (Atp7aMo-ml) causes a clinical phenotype similar to Menkes disease. Hemizygotes die by 2 weeks of age unless given supplemental copper, but female heterozygotes survive and develop an ocular phenotype. Both show 20% fewer myelinated axons than normal littermates.58 The choroid and RPE have fewer pigment granules than normal.

OPTIC NERVE DEGENERATIONS AND INFLAMMATION WALLERIAN DEGENERATION The term Wallerian degeneration refers to the changes that occur in the distal part of a cut or crushed nerve either in the peripheral or central nervous systems. C57BL/Ola mice carry a mutation in the Wallerian degeneration gene on Chr 4 that causes delayed degeneration (Wallerian degeneration slow-Wlds).45,59,60 In normal mice, following enucleation of the eye, the remaining optic nerve begins to degenerate within 5 days with myelin swelling and breakdown by 21 days after enucleation. The optic nerve in Wlds mice is normal until 21 days of age, when the degenerative process begins. The only difference in the degeneration process between normal and Wlds mice is this delay.59 The wabbler-lethal mouse (wl/wl) develops focal spontaneous Wallerian degeneration of the optic nerve as well as in the central nervous system.45 The widespread changes in the brain result in death around 1 month of age. Axons in the optic nerve are swollen and redundant myelin develops undulating folds. Both astrocytes and oligodendrocytes are normal both by light and electron microscopy.61

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AXONAL DEGENERATION Loss of axons in the optic nerve occurs in several circumstances. In glaucoma, both retinal ganglion cells and their axons die. For example, in aging DBA/2J and AKXD-28/Ty mice with advanced glaucoma, as many as 95% of normal optic nerve axons disappear (see Chapter 8).62-64 In some conditions, focal loss of retinal ganglion cells produces a corresponding focal loss of axons in the optic nerve, since axons from a specific retinal sector remain associated within the optic nerve. Focal optic nerve degeneration occurs in C57BL/6J p53–/– mice.65 Before the age of 9 months, the number of axons in the optic nerve appears constant,66 but there is a suggestion that as many as 15% of axons may be lost by 22 months of age.67 However, these results must be viewed with caution. Careful studies have demonstrated considerable variation in the numbers of retinal ganglion cells and optic nerve fibers in different strains of mice.68,69 Furthermore, accurate counting of axons in the optic nerve requires careful attention to morphometric technique (Smith and John, personal observations). While the issue of possible aging changes remains interesting, definitive quantitative studies are currently not available.

OPTIC NEURITIS Human multiple sclerosis has both a demyelinating (see above) and inflammatory component. In the optic nerve, the latter effect is frequently one of the first clinical signs of multiple sclerosis. Experimental allergic encephalomyelitis (EAE) mimics the inflammatory component. The disease can be induced by immunizing mice with optic nerve or spinal cord homogenates or with central nervous system myelin components including proteolipid protein and myelin basic protein (reviewed in O’Neill et al.70). The disease develops between 15 and 20 days after injection as an infiltration of the optic nerve (and other parts of the central nervous system) by lymphocytes and eosinophils (see Chapter 10).70,71 These cells are also found beneath the pia and in the form of perivascular infiltrates. Demyelination of the optic nerve does not occur during the acute phase. However, 35 to 40 days after immunization, there is an inflammatory relapse with additional lymphocytic infiltration. At this time, demyelination and gliosis are prominent features. Perivasculitis accompanies the inflammation.70

ORBIT DEVELOPMENTAL ABNORMALITIES Few developmental abnormalities of the orbit have been described in mice, although it is likely that subtle malformations are missed if there is no grossly visible orbital phenotype. Orbital effects have been reported in mutations affecting the isoforms of retinoic acid receptors (RARs), in addition to the anterior segment and retinal abnormalities previously discussed (see Chapters 8 through 10). Occasional agenesis of the Harderian glands occurs in nearly all of the RAR mutations. The highest incidence is in RARα1γ and RARβ2γ double null mutants.35,36,72

LACRIMAL GLAND—SJOGREN’S SYNDROME In humans, Sjogren’s syndrome is a common disorder, especially in post-menopausal women, characterized by dry mucous membranes, including those in the eye.73-75 In this autoimmune disease, there is diffuse lymphocytic infiltration of lacrimal and salivary glands that may ultimately be associated with lymphoid neoplasms (reviewed in Tsubota and colleagues76,77). Severe dry eyes may lead to secondary problems including visual loss, corneal neovascularization, and infectious corneal ulcers. Secondary forms of Sjogren’s syndrome are seen in rheumatoid arthritis, scleroderma, and systemic lupus erythematosis (reviewed in Jabs and Prendergast78). Sjogren’s syndrome has been described in

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several mouse strains including MRL/Mp-Tnfrsf6lpr, NZB/W, NFS/Njic-sld, TGF-β1 null, NODCB17-Prkdcscid, and in NOD mice that express the soluble TNF-receptor p55.77-83 Morphological changes in the affected strains demonstrate findings that include development of autoantibodies, chronic glomerulonephritis, and lymphocytic infiltration of lacrimal and salivary glands by both T and B cells (Figure 11.5).78,81 The presence of systemic disease suggests that these mice are primarily a model for lupus erythematosis with a secondary Sjogren’s syndrome.84 In NFS/Njic-sld mice, there are circulating autoantibodies and lacrimal gland lymphocytic infiltrates but no systemic disease, findings more compatible with a primary Sjogren’s syndrome.77 It is

FIGURE 11.5 Harderian and lacrimal inflammation. A. Harderian adenitis is usually focal (arrow). B. Widespread inflammation has led to extensive necrosis of the Harderian acini. For the most part, nuclei of the acinar cells have disappeared (compare to A). C. One part (*) of the Harderian gland has been destroyed by the acute inflammatory response and there is intense inflammation (arrow) at the junction between necrotic tissue and surviving gland. Original magnification × 100. D. At higher magnification, neutrophils and lymphocytes have replaced normal acinar tissue. E. Asymptomatic focal lacrimal gland inflammation (arrow) is commonly seen in some strains, such as this SWR/J mouse. This finding is especially common in older mice. Original magnification × 200. F. More widespread lacrimal adenitis is less common. Original magnification × 400.

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important to note that most, if not all, immunocompetent mouse strains, maintained under conventional housing conditions, develop various patterns of lymphocytic periductal inflammation of lacrimal and salivary glands. These changes are common in older mice and increase in severity with age. It is more likely that this aging condition is strain dependent and also reflects environmental conditions rather than being a true homologue for Sjogren’s syndrome (Sundberg and Bronson, unpublished observations).

THYROID ORBITOPATHY Thyroid eye disease is usually associated with either thyroiditis or hyperthyroidism and occurs more commonly in women. There is diffuse swelling of the extraocular muscles, mast cell and lymphocyte infiltration, fibrosis, fat deposition, and increased orbital glycosaminoglycans that lead to diffuse orbital edema. These processes within the orbit cause the eye to protrude (exophthalmos) and this produces corneal exposure and its complications.48 Thyroid eye disease seems clearly associated with autoimmunity, but the autoantigen has not been identified.85-87 Spontaneous, genetically induced thyroid disease has not been reported in mice. However, by using a complex immunization protocol, thyroid eye disease can be produced in BALB/c mice. The disease begins with thyroiditis, accompanied by both T and B cells, with development of circulating autoantibodies. Affected mice develop orbital disease with adipose tissue accumulation, edema, swelling of extraocular muscles, and infiltration by lymphocytes and mast cells (Figure 11.6).87 In contrast, NOD mice exposed to the same immunization developed thyroiditis, but no orbital disease. This is likely due to the presence of susceptibility or modifier genes (see Chapter 6).

ORBITAL INFLAMMATION Primary orbital inflammatory disease other than induced thyroid disease has not been reported in mice. In nearly all cases, orbital inflammation is secondary to infection involving the lids, conjunctiva, or cornea.88,89 The frequent occurrence of blepharoconjunctivitis in certain strains of mice has already been described in Chapters 4 and 8. It is possible for the superficial disease to spread into orbital structures in either a diffuse or focal inflammatory response (reviewed in Smith et al.90). Focal periorbital cellulitis may be caused by a foreign body and may spread to involve other orbital tissues. Orbital cellulitis may also be a consequence of a lid or sinus infection. Diffuse infiltration of the lacrimal and Harderian gland and focal abscesses may be seen (Figure 11.7).90,91 Commonly identified opportunistic organisms include Pasturella pneumotropica, Corynebacterium spp., and Lactobacillus spp. Although not exclusively true, albino strains seem more often affected by blepharoconjunctivitis than pigmented mice. This parallels the situation in human eyes, where fairskinned individuals are more prone to blepharitis.90 The term mollicutes is used to refer to a diverse group of small, cell wall–deficient and intracellular-dwelling bacteria that include various species of Mycoplasma, Spiroplasma, and Acholeplasma. When mollicutes of unknown variety were inoculated into the lids of mice, chronic orbital disease and exophthalmos developed. Morphologically, there was episcleral vasculitis, myositis, and diffuse orbital lymphocytic infiltration. Mollicutes were identified within the cytoplasm of orbital leukocytes by electron microscopy.92

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FIGURE 11.6 Autoimmune orbital thyroid disease. A and C. Normal orbital striated muscle bundles. Note the regular and compact arrangement of the individual muscle bundles. B and D. BALB/c recipients of human thyrotropin receptor-primed splenocytes. The muscle bundles have been separated by edema and a few inflammatory cells are present. A and B original magnification × 130; C and D original magnification × 320. (Illustrations courtesy of Marian Ludgate.) (From Many, M.C. et al., J. Immunol., 162:4966–4974, 1999. With permission.)

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FIGURE 11.7 Orbital inflammation. A. A focus of periorbital cellulitis (arrow) was caused by a small intraorbital foreign body. Several giant cells (arrowhead) are located adjacent to acute and chronic inflammatory cells. Original magnification × 200. B. Neutrophils and lymphocytes infiltrate orbital fat (F) and loose connective tissue. There is mild myositis, involving an extraocular muscle (M). Original magnification × 200. C. Orbital infection with Actinomyces ssp. is uncommon, but can produce severe inflammation. Large clumps of organisms (arrows) are surrounded by inflammatory cells, displacing the eye in the orbit. Original magnification × 20. D. The inflammatory infiltrate is mainly lymphocytic. Original magnification × 100.

NEOPLASMS NEUROFIBROMATOSIS The human eye and orbit are affected in many ways by neurofibromatosis. Clinical findings include neurofibromas of the lid and orbit, iris nodules, infantile glaucoma, optic nerve glioma, and optic nerve sheath meningioma.48 Although demonstrating other characteristics of neurofibromatosis, the mouse targeted mutations that are models for human neurofibromatosis 1 (Nf1) and 2 (Nf2)93-95 fail to show any ocular or orbital manifestations. Iris nodules composed of fibroblasts occur in transgenic mice that express the human T-cell lymphotropic virus type 1 (HTLV-1) tax gene. Morphologically, the iris nodules are different from those seen in human eyes, and there is no other ocular or orbital evidence of neurofibromatosis in these mice.96

LYMPHOPROLIFERATIVE DISEASES Lymphomas in human patients may develop both intraocular and orbital involvement. Intraocular infiltration by malignant lymphocytes is particularly likely to occur in primary central nervous system lymphoma, especially in immunocompromised populations.48,99 Spontaneous lymphoproliferative disease is known to affect intraocular structures in mice (see Chapter 9), particularly the uveal tract

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FIGURE 11.8 Orbital and ocular lymphoproliferative disease. A. A few neoplastic lymphocytes surround the orbital lacrimal gland. Original magnification × 200. B. The choroid (arrow) is heavily infiltrated by tumor cells. Original magnification × 400. C. Neoplastic lymphocytes surround the optic nerve (arrows) and infiltrate adjacent posterior orbital tissues. Original magnification × 100. D. There is widespread infiltration of the choroid, sclera, and orbital connective tissue. Original magnification × 400.

and retina. The same strains of mice that develop late-onset systemic lymphomas (BALB/c, SJL, DBA/2, C57BL, C58, NZB, and many transgenic strains)97,98 also acquire orbital lymphoid infiltrates (Figure 11.8). Lymphomatous infiltration of the eye, orbit, and brain of mice develop when 6- to 11day-old BALB/c mice are given an intraperitoneal inoculation of S49 mouse lymphoma cells.99 This provides a model that is morphologically quite similar to the disease in immunocompromised human patients.

LACRIMAL AND HARDERIAN Spontaneous lacrimal gland tumors in mice have not been reported. Lacrimal gland carcinomas occur in transgenic mice that carry mouse mammary tumor virus (MMTV) promoter driving the Erbb2 (formerly c-neu) oncogene.100 In contrast, proliferative lesions involving the Harderian glands are much more common. Harderian gland adenomas often occur bilaterally in older mice. Adenomas are more common in females than males and the incidence varies in different strains. Harderian gland adenomas and carcinoma are most common in BALB/c mice (80% in a series of 574 Harderian neoplasms).101 In BALB/c females, as many as 14% develop tumors by 17 months of age. These lesions begin as foci of hyperplasia and progress to cystic or papillary adenomas. Adenocarcinomas and undifferentiated carcinomas are also found in old mice (Figures 11.9 and 11.10). Regional and distant metastases have been reported, but are uncommon.100,102 A similar pattern of tumor evolution occurs in transgenic mice that carry the MMTV long terminal repeat linked to RAS family oncogenes.103,104

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MYOEPITHELIOMA Myoepithelial cells occur in salivary, lacrimal, and other simple and compound glands. These cells are modified epithelial cells capable of contracting, a function said to force secretory material into the excretory ducts (reviewed in Sundberg et al.105). Myoepithelial tumors are uncommon in human and other mammals, but are often seen in certain mouse strains. Myoepitheliomas most often occur in females and are most frequent in BALB/c and A/J strains of mice. The mean age of detection is 7 to 8 months of age.105 Nearly 75% of myoepitheliomas occur in the salivary glands, but periorbital (lacrimal), abdominal, and perineal sites have been reported. The tumors usually contain a central cystic cavity that is the result of liquefactive necrosis. Viable tumor cells are arranged in a palisading pattern and contain abundant eosinophilic cytoplasm (Figure 11.11).105

FIBRO-OSSEOUS PROLIFERATION The term fibro-osseous lesion has been used as an inclusive term for a variety of proliferative and degenerative bony lesions including osteodystrophy, hyperostosis, myelofibrosis, osteoporosis, osteofibrosis, and osteosclerosis (reviewed in Albassam amd Courtney106). Females are affected most often and frequency varies from 40 to 100% in older B6C3F1 mice. The etiology of the lesion is uncertain, although estrogen may play a role; plasma alkaline phosphatase levels are increased.106 The characteristic lesion demonstrates partial replacement of marrow cavities by fibroblastic cells, osteoclasts, and osteoblasts. In Figure 11.11, the proliferative lesion has caused one of the orbital bones to expand into the orbit.

FIGURE 11.9 Harderian gland papilloma. A. This lesion contains both papillary (arrow) and cystic (*) regions. Original magnification × 100. B. A small lobule of normal Harderian gland contains porphyrin deposits (arrow). Original magnification × 50. C. At low magnification, it appears that proliferating cells are less organized than in A and B. Original magnification × 50. D. At higher magnification, it is clear that the proliferating cells are well differentiated and cytologically benign. Original magnification × 200.

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FIGURE 11.10 Malignant orbital tumors. A. In contrast to Figure 11.8, this Harderian adenocarcinoma is highly cellular, and rather than producing acinar structures, the cells are growing in cords (arrows). Original magnification × 100. B. At higher magnification, the cells are completely undifferentiated, characterized by small densely staining nuclei and scanty cytoplasm (arrow). Numerous foci of tumor cell necrosis are present (*). Original magnification × 200. C. This Harderian adenocarcinoma is more differentiated, containing pseudocystic structures (*). The normal gland (arrow) is compressed and displaced by the neoplasm. Original magnification × 200. D. At higher magnification, the pseudocystic areas are lined by poorly differentiated acinar cells. Undifferentiated cells (arrow) are also present within the neoplasm. Original magnification × 200. E. An extensive squamous cell carcinoma (arrow) has invaded the orbit, compressing adjacent tissues. Original magnification × 100. F. Despite the aggressive growth of the neoplasm (E), the individual cells are well differentiated. However, mitotic activity (arrows) is frequent. Original magnification × 400.

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FIGURE 11.11 Orbital tumors. A. Orbital myoepithelioma in a 10-month-old BALB/c mouse. These are highly cellular tumors that frequently contain a central cystic cavity (*). Original magnification × 50. B. The stroma of myoepitheliomas is scanty (arrow), and the limited blood supply likely contributes to the central necrosis (A). Original magnification × 200. C. The individual myoepithelioma cells have round to oval vescicular nuclei and abundant eosinophilic cytoplasm. The tumor cells show some tendency toward palisading. Original magnification × 400. D. A large region of fibro-osseous proliferation has caused an orbital bone (arrow) to extend into the orbit, nearly impinging on the eye (E). The marrow has been replaced by an expanding lesion consisting of fibrous tissue and cells (*). Original magnification × 100. E. Higher magnification demonstrates the intimate relationship of normal bone (B) to the fibrous (F) and cellular (C) components. The cellular elements show a prominent palisading pattern. Original magnification × 400. F. Close to the normal bone (B), the cellular morphology (arrow) resembles that of osteoblasts. Original magnification × 400.

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Section IV Biomethods

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Photography and Necropsy Richard S. Smith, Norman L. Hawes, James Miller, John P. Sundberg, and Simon W. M. John

CONTENTS Photography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 251 Gross Photography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .251 External Ocular Photography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .252 Photographic Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .254 Fundus Photography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .256 Fluorescein Angiography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .256 Skeletal Preparations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .257 Radiographic Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .258 Necropsy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .258 Necropsy of the Head . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .258 Eyelids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .258 Lid and Orbital Glands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .259 Skull and Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .261 Tissue Trimming . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .262 Enucleation of the Eye . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .262 Optic Nerve—Pulling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .263 Optic Nerve—Dissection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .263 Optic Nerve—Brain Dissection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .264 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .264

PHOTOGRAPHY Phenotypic description is an important step in the process of characterization of spontaneous and induced mutant mice.1,2 Documentation of ocular clinical and necropsy findings contributes to determination of the phenotype, and for that reason these subjects are considered together in this chapter. Although subsequent discussion is concentrated on the eye, a thorough necropsy is necessary when evaluating the effects of any new mutation. Genes that affect ocular structures may also have systemic effects. Conversely, systemic diseases can produce secondary ocular effects. Failure to evaluate all organ systems can result in failure to fully comprehend important disease mechanisms. The techniques for performing complete necropsies are detailed elsewhere,3 and discussion of necropsy techniques in this chapter is limited to methods for assessment of the eye, eyelids, orbit, and brain.

GROSS PHOTOGRAPHY Even in the mouse, documentation of many clinical features before or during necropsy can often be accomplished by using a 35-mm single-lens reflex camera system with a macrolens mounted on a copy stand.4 However, even with extension tubes, the small size of the mouse eye severely limits the value of this technique. The short distance between camera lens and eye needed to achieve adequate 0-8493-0864-X/02/$0.00+$1.50 © 2002 by CRC Press LLC

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magnification makes proper illumination for photography difficult. A better arrangement is to utilize a dissecting photomicroscope with two or more fiber-optic illuminators (Figure 12.1). A foot pedalactivated camera shutter allows restraint of the mouse by the operator and production of external photographs. It is also possible to view the fundus by applying a microscope coverslip lubricated with methylcellulose to the corneal surface. A slit lamp with integrated photographic unit (Figure 12.2) provides the highest quality detailed photographs of the anterior portion of the eye in living mice (lids, cornea, conjunctiva, iris, and lens) without the need for anesthesia.5,6 If at all possible, anesthesia should be avoided because of the risks of corneal drying and the potential for occurrence of a transient anesthesia-related lens opacity.7 High-quality photographs of the retina, choroid, and optic nerve can be produced by using a small-animal fundus camera (Figure 12.3).8 Together, these two techniques are superior replacements for the traditional gross photographs of the general pathologist.

EXTERNAL OCULAR PHOTOGRAPHY For those unfamiliar with the slit lamp, the older term “biomicroscope” suggests its function. A slit lamp has an illumination source that can be narrowed to a fine linear beam. Combined with the attached low-power microscope (5 to 60× magnification), it is possible to create an optical “section” of the area being viewed (Figure 12.4). With appropriate manipulation of the slit beam and

video

35mm camera

fiber optic light

FIGURE 12.1 Photodissecting microscope. Variable-focus dissecting microscope equipped with dual fiberoptic lights and adapters for video and 35-mm camera (Olympus).

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camera

platform

FIGURE 12.2 Slit lamp. Typical slit lamp with 35-mm camera attachment (Haag–Streit, Switzerland) adapted for mouse photography. The chin rest is replaced by a cleanable Plexiglas platform that helps steady the mouse for photography. Attachment of a video system is also possible.

microscope, individual corneal endothelial cells can be visualized, as well as details of the corneal stroma, anterior chamber, iris, and lens structure. Illumination with the broad beam provides a full view of the anterior ocular structures. Different photographic attachments can be integrated with the basic slit lamp including digital computer image capture devices or standard 35-mm cameras. The latter remain the best source of publication quality pictures, although digital technology will likely replace older methods in the near future. Integrated flash units are needed to obtain short exposure times. The flash system should be equipped with adjustable flash intensities and apertures as well as a rapidly recharging power system. Different intensities are critical for mice with various degrees of ocular pigmentation. An exposure suitable for a black mouse will yield a washed-out image from an albino animal. Since systems vary, a few calibration rolls of film are usually needed before a routine is established.

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FIGURE 12.3 Small animal fundus camera (Kowa, Tokyo, Japan) adapted for mouse fundus photography (power supply not shown). The camera is supported by a dissecting microscope stand that allows for coarse focus adjustment. The condensing lens is supported by a holder that is attached to a magnetic stand. This arrangement frees the operator to position the mouse and focus the camera by slight movements of the mouse.

PHOTOGRAPHIC TECHNIQUES Mouse slit lamp photography is easier if the chin rest designed for human use is removed and replaced with a clear plastic platform (see Figure 12.2). The mouse is gently restrained with one hand resting on the platform for stability. The other hand can be used to retract the lids and provide additional stability. The platform also protects the equipment from feces and urine. The shutter is activated by using a foot pedal. Although the slit lamp is normally focused by moving it on a sliding track when human patients are examined, it is easier to lock the optical system in place and move the mouse to focus on the eye. These techniques provide a clear view of the entire anterior segment of the eye for clinical phenotyping or photography and permit efficient screening of large numbers of mice. Broad-beam illumination is best in most situations. The slit beam is useful for localizing the depth of vessels or opacities in the cornea or for determining the extent and location of partial lens opacities. A magnification of 16 to 25× is suitable for screening purposes. A magnification of 40× is ideal for photodocumentation since the eye nearly fills the 35-mm image field and fine details of ocular structures are easily resolved (Figure 12.5).

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FIGURE 12.4 Slit lamp function.When the slit lamp is adjusted to form a thin rectangle of light, an optic section of the eye is produced. This allows visualization of details of the anterior corneal epithelial surface (A), the corneal stroma (S), and the posterior endothelial (E) surface. Lens and iris structural details may be identified. A deeper focus (not shown) will provide an optical section of the lens.

FIGURE 12.5 Examples of broad-beam slit lamp photographs of pigmented and albino mouse eyes. A. In a 21month-old B6D2F2 mouse there is severe atrophy of the iris sphincter (arrowhead) and diffuse iris stromal atrophy (*). Even with the broad beam, peripheral iris transillumination (arrow) is easily visualized. B. A 9-month-old 129/SvJ albino mouse demonstrates superficial corneal neovascularization (arrowhead). Fine iris vessels (arrow) and details of iris structure are demonstrated. This photograph was taken with illumination intensity approximately one half that used to produce A.

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FUNDUS PHOTOGRAPHY Phenotypes that produce changes in the retina, optic nerve, and vitreous can be recorded using a fundus camera. Although human fundus cameras have been used for this purpose, the small size of the mouse eye and the relatively small pupil make it difficult to obtain high-quality fundus photographs consistently. In our hands, better results are obtained by using a camera (see Figure 12.3), such as the Kowa Genesis, specifically designed for small-animal fundus photography (Kowa, Tokyo, Japan).8 A condensing lens (60 to 90 diopter) mounted between the camera and eye provides adjustable magnification and allows the photographer to restrain the mouse gently as described for slit lamp photography (for details, see Hawes et al.8). It is necessary to dilate the pupils 5 to 10 min before photography. A single drop of 1% atropine or 1% cyclopentolate is usually sufficient to achieve adequate dilation, although some mice are more resistant to the drops than others.8 Different parts of the fundus can be photographed by changing the mouse’s position relative to the condensing lens. Focusing is critical because the small diameter of the mouse eye and the consequent steep radius of ocular curvature give little depth of focus. The flash unit provided with the camera has adjustable settings that can be changed depending on fundus pigmentation (Figure 12.6).8

FLUORESCEIN ANGIOGRAPHY This technique is utilized for demonstrating vascular structures in the eye, particularly in the retina, choroid, and optic nerve. Typical features detectable on an angiogram include circulation through the iris vessels; arterial, capillary, and venous circulation of the retina; choroidal and superficial optic nerve vascular systems; vascular leakage; and abnormal angiogenesis. When fluorescein angiography is performed in human patients, a bolus of dye is injected intravenously and the circulation of the dye through the ocular vascular systems is followed by rapid sequence photography. The dye appears in the eye in 10 to 12 s, appearing first in the arterioles. Vascular leakage is detected by taking photographs 20 s to 30 min after injection, as the extravasated dye usually remains and there is often progressive leakage.

FIGURE 12.6 Normal pigmented and albino fundi from the right eye. A. C57BL/6J mouse, 8 weeks of age. Venules (V) are twice the diameter of arterioles (A). The arteriolar blood column has a copper tint and is more reflective, while the blood in the venule is dull red. B. BALB/cByJ mouse, 12 weeks of age. The choroidal vessels are visible through the retina because of absence of choroidal and retinal pigment epithelial pigmentation. The optic nerve head is indicated by an arrow and the central retina by an asterisk. (From Hawes, N.L. et al., Mol. Vision, 5:22–29, 1999. With permission.)

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Although mice can be given intravenous fluorescein, anesthesia is usually required and placement of the needle is technically difficult. Angiography can be performed in unanesthetized mice by intraperitoneal injection of small amounts of fluorescein. Satisfactory results are obtained with 0.01 ml per 5 to 6 g body weight of 25% sodium fluorescein (Akorn, Inc., Decatur, IL). For photography, the standard fundus camera back is replaced with one containing a barrier filter and the power pack is set for angiography, which places barrier and exciter filters in the appropriate position. Direct viewing is also possible by fitting the eyepiece with a manufacturer-supplied barrier filter.8,9 With intraperitoneal injection, the first intravascular dye is detected in about 30 s and complete washout occurs by 5 min. When present, vascular leakage has usually been detected 1 to 5 min after injection (if leakage occurs, the extravascular dye persists for longer periods) (Figure 12.7).8

SKELETAL PREPARATIONS Bones surrounding the eye can vary between strains10,11 or be very abnormal in some diseases. The skull can be evaluated by removing most of the soft tissues, clearing those remaining with 0.5 to 2.0% potassium hydroxide, and staining the bones with alizarin red S, which aids visualization of skeletal details (Figure 12.8).12 The noncleared skulls illustrated in Figures 2.7 and 2.8 were prepared by 1% potassium hydroxide digestion over a period of 1 week with changes to fresh solution every 2 days and periodic manual removal of tissue fragments. Final cleanup was done by boiling over low heat for 1 h. FIGURE 12.7 Angiography. Fluorescein angiogram from an 8-week-old C57BL/6J mouse, in early venous phase. The retinal arterioles (A) are smaller than the venules (V). The complex retinal capillary bed (C) is clearly demonstrated.

FIGURE 12.8 Cleared skulls stained with alizarin red S illustrate normal C57BL/6J (A) and very thin (B) zygomatic arches (arrows), the latter in the defective hair and ears (Dhe/+) mutant.

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RADIOGRAPHIC TECHNIQUES A small X-ray machine (Faxitron, Hewlett-Packard model 43855A), produced to evaluate electronic boards, provides a simple and rapid way to evaluate bones of the mouse. The mouse must be anesthetized or euthanized to prevent movement during exposure. Another alternative is the MX-20 specimen radiography device (Faxitron X-ray, Wheeling, IL) which produces very high resolution radiographs that can demonstrate subtle bone structure changes and detect small foci of intraocular mineralization. Lateral and dorsoventral views are taken using mammography film (Kodak X-O Mat TL-Eastman Kodak, Rochester, NY). The 8 × 10 in. sheets of film are enclosed in a paper envelope to prevent exposure to room light. Multiple images can be taken on the same sheet of film by using lead plates to cover all but the area being exposed. The outside of the film packet can be marked to distinguish between exposed and unexposed portions of the film. Lead letters and labels (TechnoAide, Nashville, TN) are placed on the edge of each frame so they do not block view of the skull. The skull orientation is indicated by the letters left (L) and right (R) side, dorsoventral (DV) or ventrodorsal (VD) orientations, and additional letters and numbers are used to identify the specimen (accession number, date, etc.). These provide permanent identification of each image for easy record keeping. Exposed film is processed routinely in an automated X-ray film processor or by hand in standard chemical tanks with metal holders for X-ray film of the size being used. Type 55 Polaroid film (Polaroid Corp., Cambridge, MA) provides instant results, with a positive print and a negative. Unfortunately, resolution is inferior to mammography film. These methods can be used to evaluate subtle to dramatic changes in small bones such as the zygomatic arches around the eye (Figure12.9). A major problem with radiographs is evaluating overlapping structures, since the image superimposes the left and right structures in lateral views and the maxilla and mandible in dorsoventral views. The mandible is removed from the skull for a dorsoventral view. This provides left-right symmetry. The skull is bisected on the midline suture of the calvarium using a sharp, single-edged razor blade. Both halves are placed cut side down, labeled L and R, respectively, and the lateral radiograph taken. This also works well for evaluating teeth.11 Computer scanning of radiographs provides a simple method to provide copy for lecture slides or publication (Figure 12.9).

NECROPSY NECROPSY OF THE HEAD There are two approaches useful for preparing the eyes and adnexae for histopathology: examination in situ and enucleation. The former is utilized for routine screening purposes as well as being the method of choice for mice under 2 weeks of age. In young mice and embryos, decalcification of the skull is not needed if the head is fixed in Bouin’s solution (see Chapter 15) and carefully oriented in the histology cassette. Eyes from older mice are best removed and processed separately (see below). Eyelids For comparison of adult mice, use of standardized methods for preparing the head for sectioning ensures consistent results that can be used for comparison. To evaluate the eyelids histologically, there are two approaches. The eyelids can be removed by cutting the skin around the orbits and carefully dissecting them free using a scalpel with gentle traction by iris forceps. Alternatively, all skin of the head, including the eyelids, is carefully peeled and trimmed from the skull as a unit and mounted flat on a piece of foil for fixation (Figure 12.10). Following fixation, the eyelids are trimmed to assure proper orientation for histology.3 Eyelids are usually presented by using a razor blade to cut vertically through the center of both lids. A second cut is made parallel and 4 to 5 mm from the first, and should include enough facial skin to keep the eyelids together (Figure 12.11). The face of the second cut is

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FIGURE 12.9 Radiographs of C57BL/6J+/+ (A) and Dhe/+ (B) mice illustrate marked reduction in bone size of the zygomatic arch (white arrows).

marked with blue pencil (Venus col-erase, #1276 blue, Eberhard Faber, Inc., Lewisburg, TN) to help with orientation. Lid and Orbital Glands The Meibomian glands are usually well oriented in standard eyelid preparations. Longitudinal trimming (along the length of the eyelid margin) will provide a perspective of gland distribution. The lacrimal and Harderian glands are usually found in coronal sections of the skull that include the eyes. Each gland can also be dissected individually. The Harderian gland is a large, soft, lobulated, light brown structure that is easily identified and removed after enucleation of the eye. In mice, the

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FIGURE 12.10 Skin orientation. A piece of skin has been flattened on aluminum foil to prevent rolling and wrinkling in fixative. A similar technique can be used to assist in orienting the eyelids.

FIGURE 12.11 Head skin trimmed for histology. The indicated razor blade cuts create three separate blocks of tissue that include the muzzle and vibrissae (A), the eyelids, cilia, Meibomian glands, and conjunctiva (B), and the pinna of the ear (C). (Courtesy of Ingrid Sundberg.)

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lacrimal gland is divided into two separate portions that empty into the conjunctival sac via a common duct. The orbital part of the lacrimal gland is located laterally beneath the upper lid; it is best visualized in the coronal skull section. The extraorbital portion of the gland is positioned anterior and ventral to the ear, adjacent to the parotid gland. The nictitating membrane, located by the medial junction between upper and lower eyelids may sometimes be identified on the coronal skull section, but is often missed because of its small size. Skull and Brain The skull is processed separately. All skin is removed, including the eyelids. At the foramen magnum, one blade of the scissors is slipped between brain and bone and two small longitudinal cuts are make through the occipital bone to the lambdoid suture (Figure 12.12), one on each side of the spinal medulla. The skull is held between the thumb and forefinger, and with forceps the edge of the occipital bone is lifted upward and removed. The tip of the scissor blade is inserted between the brain and skull on the sagittal suture and a cut is made a short distance past the coronal suture (Figure 12.13). The interparietal and parietal bones are gently broken along the cut edge to increase exposure of the brain. The frontal bones come to a slight point at the intersection between sagittal and coronal sutures. This point must be removed to allow removal of the brain without damage. After removal of the meninges, forceps are used to loosen the brain from the skull by gently pulling away residual connective tissue and the cranial nerves still attached to the brain. This frees the brain, which is placed into fixative by inverting the skull.

FIGURE 12.12 Skull necropsy technique. Two parallel cuts are made in the occipital bone to expose the brain. (Courtesy of Ingrid Sundberg.)

FIGURE 12.13 Skull necropsy technique. A midline cut through the sagittal suture and removal of the bone on either side of the cut increases exposure of the brain. (Courtesy of Ingrid Sundberg.)

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TISSUE TRIMMING Trimmed tissues that were fixed in formalin-based fixatives and not decalcified can be placed in histology cassettes for storage in 70% ethanol. Tissues that require decalcification (demineralization) should be placed in a container with 1.35 N hydrochloric acid (CAL-EX, Fisher Scientific, Pittsburgh, PA) and held overnight at room temperature. A longer exposure will significantly reduce eosin staining. After decalcification, the tissues are rinsed in running tap water in histology cassettes for a minimum of 12 h. After rinsing, decalcified tissues are placed in 70% ethanol. All non-eye tissues should be trimmed to 1 to 2 mm in thickness for optimal penetration of solvents and paraffin. The skull is trimmed as three cross sections for routine examination (Figure 12.14) to include the eyes, nasal passages, ears, and pituitary gland. The first razor blade cut should be made immediately posterior to the edge of the pituitary, which can be identified as a small pink or white mass that lies sagitally on the floor of the skull between the right and left trigeminal ganglia (two thick nerves with a longitudinal course). The second cut is made 4 to 5 mm anterior to the first cut to create the first section. In addition to the pituitary, this tissue block includes external, middle, and inner ear structures. The third cut is made through the posterior edge of the visible portion of the eyes. The fourth cut is made 4 to 5 mm anterior to the third cut and includes the eyes and orbital structures. The final cuts are through the approximate center of the remaining portion of the snout and will include the nasal cavity. All trimmed tissues are placed in plastic histology cassettes in 70% ethanol for submission. Hematoxylin and Eosin (H&E) stain is routinely requested on all tissues sent to the histology laboratory. Staining protocols and their appropriate applications are widely available13,14 and are discussed in detail in Chapter 13.

FIGURE 12.14 Skull necropsy technique. Three sections are usually cut in the decalcified skull. The dotted lines indicate the width and locations of the tissue cuts. These cuts expose the nasal cavity (A), the eyes and orbital structures (B), and the inner, middle, and external ear, as well as the pituitary gland (C). (Courtesy of Ingrid Sundberg.)

ENUCLEATION OF THE EYE Careful enucleation and processing of eyes from adult mice is critical for producing sections that are reliable and interpretable. Eyes must be removed immediately following euthanasia to avoid postmortem artifacts. The easiest enucleation technique is to place a curved serrated forceps well behind the globe, grasp the tissue firmly, and gently pull forward. Grasping directly behind the globe is undesirable, because the optic nerve may be crushed and the contents squeezed into the eye. This results in what is termed a myelin artifact, because the myelin of the nerve is displaced. (Figure 12.15)

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FIGURE 12.15 Myelin artifact. Retina and optic nerve from a 6-month-old DBA/2J mouse. The retina (R), retinal vessels (V), and optic nerve head (N) are well preserved. Because this nerve was cut too close to the globe, the myelin from the optic nerve was extruded from the nerve to a location between retina and choroid (region between asterisks).

Placement of the forceps must be done with care to avoid perforation of the globe or blunt trauma, which can cause intraocular bleeding and retinal detachment. When this technique is used, the optic nerve, especially in older mice, may separate at its juncture with the globe. Overall, pulling is the most satisfactory technique and can produce long excellent sections of optic nerve. However, if there is optic nerve disease, as occurs in older DBA/2J mice, crucial damage of the nerve may occur. Alternate methods are needed to avoid these problems. Optic Nerve—Pulling Several approaches can be employed to obtain a long, undamaged section of optic nerve. In younger mice (less than 4 months of age), appropriate use of curved forceps and gentle traction is usually satisfactory. If the caudal and inferior exit of the optic nerve from the orbit (Chapter 4) is kept in mind, the forceps can be placed gently behind the globe and pushed in that direction as far as possible, with the tips open. The optic nerve can be grasped at the end of this maneuver and several millimeters of undamaged nerve obtained. Optic Nerve—Dissection A second approach is to expose the nerve by a combination of sharp and blunt dissection. The limbal conjunctiva is cut for 360°, close to the corneoscleral junction. Because of the orbital venous sinus, this often produces considerable bleeding, even in a euthanized mouse. This ceases fairly rapidly and then the caudal portion of the upper and lower lids can be excised, followed by careful removal of the caudal portion of the Harderian gland. Care must be taken during these steps to avoid traction on the globe, which could damage the optic nerve. These procedures open enough space in the orbit to allow placement of a fine curved iris scissors close to the optic foramen, where the optic nerve may be severed. Careful attention to cutting any remaining extraocular muscles and orbital connective tissue allows removal of an intraorbital optic nerve of maximum length. In our hands, these procedures generally produce satisfactory tissue for most experimental purposes.

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Optic Nerve—Brain Dissection Older mice, especially those with glaucoma and secondary optic nerve abnormalities, present additional problems. A diseased nerve is extremely fragile and may be less than 25% normal diameter. Even careful dissection can sever the nerve close to the globe. For ideal nerve fixation, the posterior dorsal skull should be removed immediately after euthanasia. The dorsal 80% of the brain is gently removed by horizontal dissection with a sharp, disposable blade. The remaining brain tissue protects the optic nerves and chiasm from stretching that can occur if the whole brain is removed prior to fixation. After dissection, the skull is immersed in a glutaraldehyde-paraformaldehyde fixative15 for 24 h and may be stored in buffer. At the time of dissection, remaining brain tissue is removed and the optic nerves and chiasm exposed and removed. Orientation of nerve direction is achieved by leaving a portion of the chiasm attached, dipping one end in eosin, or cutting one end on a slant. An alternative approach specific to optic nerve collection is discussed in the histology section of the next chapter. Although the nerve tissue quality is superior with this procedure, the eyes become brittle when fixed in the skull. Frequently, when these eyes are removed by dissection the intraocular structures are damaged. For this reason, careful dissection and removal of the eye before fixing the whole skull is likely the best approach. Clearly, these more laborious techniques require fine dissecting instruments, patience, and practice.

REFERENCES 1. Martin, J.E. and Fisher, E.M.C., Phenotypic analysis-making the most of your mouse, Trends Genet., 13:254, 1997. 2. Beck, J.A. et al., Genealogies of mouse inbred strains, Nat. Genet., 24:23, 2000. 3. Relyea, M.J. et al., Necropsy methods for laboratory mice: biological characterization of a new mutation, in Systematic Approach to Evaluation of Mouse Mutations, Sundberg, J.P. and Boggess, D., Eds., CRC Press, Boca Raton, FL, 1999, 57. 4. Sundberg, J.P. and Miller, J., Photography of laboratory mice, in Systematic Approach to Evaluation of Mouse Mutations, Sundberg, J.P. and Boggess, D., Eds., CRC Press, Boca Raton, FL, 1999, 91. 5. John, S.W.M.J. et al., Essential iris atrophy, pigment dispersion and glaucoma in DBA/2J mice, Invest. Ophthalmol. Vis. Sci., 39:951, 1998. 6. Chang, B. et al., Interacting loci cause severe iris atrophy and glaucoma in DBA/2J mice, Nat. Genet., 21:405, 1999. 7. Calderone, L., Grimes, P., and Shalev, M., Acute reversible cataract induced by xylazine and by ketamine-xylazine anesthesia in rats and mice, Exp. Eye Res., 42:331, 1986. 8. Hawes, N.L. et al., Mouse fundus photography and angiography: a catalogue of normal and mutant phenotypes, Mol. Vision, 5:22, 1999. 9. Okamoto, N. et al., Transgenic mice with increased expression of vascular endothelial growth factor in the retina, Am. J. Pathol., 151:281, 1997. 10. Beamer, W.G. et al., Genetic variability in adult bone density among inbred strains of mice, Bone, 18:397, 1996. 11. Mahler, M. et al., Pathology of the gastrointestinal tract of genetically-engineered and spontaneous mutant mice, in Pathology of Genetically Engineered Mice, Ward J. et al., Eds., Iowa State University Press, Ames, 2000. 12. Selby, P.B., A rapid method for preparing high quality alizarin stained skeletons of adult mice, Stain Technol., 62:143, 1987. 13. Luna, L.G., Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology, McGraw-Hill, New York, 1960. 14. Smith, A. and Bruton, J., Color Atlas of Histologic Staining Techniques, Year Book Medical Publishers, Chicago, 1977. 15. Smith, R.S., Ultrastructural studies of the blood-aqueous Barrier. I. Transport of an electron-dense tracer in the iris and ciliary body of the mouse, Am. J. Ophthalmol., 71:1066, 1971.

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General and Special Histopathology Richard S. Smith, Adriana Zabeleta, Simon W. M. John, Lesley S. Bechtold, Sakae Ikeda, Melissa J. Relyea, John P. Sundberg, Winston W.-Y. Kao, and Chia-Yang Liu

CONTENTS A. Light Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 266 Richard S. Smith, Adriana Zabeleta, and Simon W. M. John Fixation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .266 Tissue Processing for Paraffin Embedding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .267 Paraffin Sectioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .267 Fixation and Embedding for in Situ Hybridization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .268 Tissue Processing for Plastic Embedding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .268 Plastic Sectioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .268 Section Interpretation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .269 Collection of Optic Nerve . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .269 Embedding and Sectioning Optic Nerve . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .269 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .272 B. Electron Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .272 Lesley S. Bechtold and Richard S. Smith Fixatives and Buffers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .272 Fixation and Tissue Trimming . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .273 Postfixation and Embedding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .274 Routine Protocols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .275 Fixation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .275 Dehydration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .276 Infiltration and Embedding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .276 Solution Preparation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .276 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .276 Suggested Reading . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .277 C. Immunohistochemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .277 Sakae Ikeda, Melissa J. Relyea, and John P. Sundberg Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .277 Primary Antibodies and Fixation Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .277 Enzyme and Fluorescence Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .278 Enzyme Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .278 Fluorescence Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .280 Overview of the Staining Procedure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .280 Tissue Preparation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .280 Incubation with Antibodies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .281 0-8493-0864-X/02/$0.00+$1.50 © 2002 by CRC Press LLC

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Enzyme Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .281 Fluorescence Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .282 General Protocols for Immunohistochemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .282 Interpretation of Results . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .283 Positive Controls . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .283 Negative Controls . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .283 Autofluorescence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .283 Endogenous Enzyme Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .284 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .284 D. Northern and in Situ Hybridization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .285 Sakae Ikeda, Chia-Yang Liu, and Winston W.-Y. Kao Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .285 Precautions against Ribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .285 Northern Hybridization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .285 Total RNA Preparation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .286 Protocols for Northern Hybridization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .286 In Situ Hybridization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .290 Fixation and Permeability of Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .290 Probe Labeling: Radioactive or Nonradioactive . . . . . . . . . . . . . . . . . . . . . . . . . . . .290 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .296 Careful attention to tissue collection, fixation, sectioning, and staining are all critical in producing slides for microscopic examination that accurately demonstrate what is happening in the tissue. Tissues such as liver and kidney are relatively easy to section, but eyes are not. The hardness of the lens as well as its delicate suspension by the zonules makes it a particular challenge. The lens shatters easily and, if infiltration or embedding is suboptimal, may move during sectioning and damage other ocular tissues. The relative thinness of the cornea, sclera, choroid, and retina make it easy for folds to develop as sections are collected on slides. Procedures for collection of eyes are discussed in Chapter 12, and Section 13A is primarily concerned with techniques of fixation, embedding, and sectioning of tissues for light microscopy. The one exception is collection of the optic nerve, which is described in this chapter, in conjunction with special sectioning and staining techniques for nerve. Special staining techniques are reviewed elsewhere in great detail,1 and will not be discussed here. The basic techniques for electron microscopy, immunohistochemistry, and in situ procedures are reviewed in subsequent sections of this chapter.

A. LIGHT MICROSCOPY Richard S. Smith, Adriana Zabeleta, and Simon W. M. John

FIXATION Both immediate and potential needs should be considered when selecting a fixative. For example, the specific requirements for an immunohistochemical procedure may eliminate fixatives entirely suitable for other purposes. Whatever the fixative, tissue should be immersed immediately after removal to avoid drying and postmortem artifacts. Protocols that describe the composition and preparation of fixatives are described elsewhere.2 Two of the more commonly used fixatives produce less than optimal results. Bouin’s solution in adult mice causes loss of fine cellular details and overstaining by eosin. However, the advantages of Bouin’s solution in fixation of the head in young mice (see below) outweigh the negative features because it facilitates sectioning by decalcification as a result of the picric acid in the fixative. Neutral 10% buffered formalin often hardens the lens, making it more likely

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to shatter when sectioned. These problems are alleviated by using Fekete’s acid-alcohol-formalin for paraffin embedding and some immunohistochemistry. Tissue is fixed for 24 h at room temperature and can be stored in 70% ethanol. Although paraffin sections are adequate for many purposes, plastic-embedded material is needed for interpretation of subtle details and for publication-quality pictures. For eye tissues, the most consistent fixation results for plastic embedding are achieved by using a 0.1 M phosphate-buffered glutaraldehyde paraformaldehyde mixture.3 Tissues are fixed for 24 h at 4°C in the dark and can be stored in 0.1 M phosphate buffer, pH 7.4 at 4°C. In addition, the same fixative can be used for tissue prepared for transmission electron microscopy (see Section 13B). An alternative fixative is 4% paraformaldehyde, which can be used for immunohistochemistry, in situ paraffin-embedded tissue, and for plastic embedding.4,5 Tissues are fixed at room temperature for 1 to 6 h and can be stored in 0.4% paraformaldehyde for at least 1 year without deterioration. Both concentrations are prepared in 0.1 M phosphate buffer, pH 7.2. The small size of the perinatal eye makes it difficult to enucleate without damage. Despite objections to using Bouin’s solution in adults, this fixative offers several advantages in mice less than 2 weeks of age. Decalcification before sectioning is not needed at these ages and potential trauma associated with enucleation of a small eye is avoided. The whole head is placed in Bouin’s solution for 24 to 48 h. The nose and back of the head are removed after this period and the remaining tissues washed briefly in running water and transferred to 70% ethanol. The ethanol is changed several times, which helps eliminate the excess of Bouin’s solution. After this, the heads are processed for paraffin embedding as described below. When the mice are 7 to 14 days of age, it is best to hemisect the skull along the midline and separately section each eye.

TISSUE PROCESSING FOR PARAFFIN EMBEDDING Eyes are dehydrated in graded alcohols at room temperature (65% once, 80% twice, 95% once, 100% three times; 1 h for each change of solution). This is followed by three 1-h changes of xylene (room temperature) and four changes of paraffin at 60°C under 15 mmHg of vacuum to remove air bubbles. During embedding, the eyes are oriented so that a horizontal plane extending through the cornea and optic nerve is as parallel as possible with the bottom of the mold.

PARAFFIN SECTIONING For the highest-quality morphology, plastic sections are always superior to paraffin. However, careful attention to technique allows production of good-quality paraffin sections that are useful for screening purposes and necessary for some techniques such as immunohistochemistry. Chilling the paraffin block and soaking it after initial rough cutting is particularly helpful. Prior to sectioning, 3 to 4 drops of concentrated (28%) ammonium hydroxide is applied to the surface of an ice cube and the paraffin block is placed face down on the ice for 15 to 30 min. An alternative is to soak a laboratory wipe with 10% ammonium hydroxide and leave the tissue blocks in contact with this solution overnight at 4°C. This maneuver softens the lens and decreases the risk of shattering. However, ammonium hydroxide cannot be used with in situ hybridization techniques. All procedures involving ammonium hydroxide should be done with adequate ventilation in a hood and with eye protection. Once prepared, the block is placed in the microtome chuck with the cornea and optic nerve at the sides of the block, in contrast to the procedure used when sectioning plastic blocks (see below). The lens frequently has a strong tendency to become detached from the slide. Addition of a single drop of Elmer’s® glue (Borden, Inc., Columbus, OH) or poly-L-lysine to the water bath will minimize the problem. Rough handling during the staining procedure can also cause lens detachment from the slide.

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FIXATION AND EMBEDDING IN SITU HYBRIDIZATION Tissues are fixed 6 h in 4% paraformaldehyde in 0.1 M phosphate buffer, followed by processing in graded alcohols as described for paraffin sectioning except that the alcohols are made with diethylpyrocarbonate (DEPC) water (Sigma Chemical Co., St. Louis, MO) After embedding (as for paraffin) the blocks are stored at 4°C. DEPC water is used in the water bath when sectioning. Sections (4 to 5 µm) are placed on glass slides and stored at 4°C with silica gel desiccant until used. The slides are rehydrated using DEPC water alcohols and treated as described elsewhere (Chapter 13D).6,7

TISSUE PROCESSING FOR PLASTIC EMBEDDING Tissues are dehydrated in 25, 50, 70%, and two changes of 95% alcohol for 1 h each, on a shaker set at high speed. The alcohol is replaced with a degassed 1:1 historesin–95% ethanol solution for 24 h. This is followed by 24 h in 100% historesin (Leica), then 3 days in a fresh change of resin; at all times a shaker is used to aid in the infiltration process. After the final historesin change, tissues are infiltrated and embedded in degassed historesin at 4°C. The embedding process can be extended to 10 days and somewhat better overall quality was achieved with the longer embedding time. The eyes are immersed in fresh embedding medium and oriented in a mold under a dissecting microscope. The mold is filled to within 2 to 3 mm of its top. It is important to position the eye so that a plane extending through the cornea and optic nerve is as parallel as possible with the bottom of the mold. Careful positioning facilitates proper orientation when sectioning begins. A dozen eyes can be embedded in a total of 16 ml of embedding medium. The rate of polymerization is rapid enough to maintain proper orientation. The process of polymerization requires at least 12 h.

PLASTIC SECTIONING The historesin tends to absorb water and this can interfere with sectioning. For this reason, it is best to leave the plastic blocks in a desiccator for 24 h prior to sectioning. When the block is sticky and sections have a tendency to stretch and wrinkle, this indicates the presence of residual water in the block. If this occurs, the blocks should be put in a 45°C incubator for 30 min and allowed to cool in a desiccator. Sectioning artifacts of the cornea can be avoided by orienting the plastic block in the microtome with the optic nerve at the bottom and the cornea at the top. To avoid damage to the block, initial trimming cuts should be no greater than a maximum thickness of 3 µm. For most purposes, sections are cut at 1.5 µm. No matter what the embedding material, the lens is the greatest barrier to section quality because of its hardness and tendency to shatter. If this problem occurs it may be helped by cutting several sections at a thickness of 0.5 to 1.0 µm. Some histologists prefer tungsten carbide knives because they produce better lens sections than glass knives. However, we feel that the overall quality of sections may be better with glass knives. We use a glass knife for rough trimming and then replace it with a fresh glass knife prior to section collection. Glass knives may lose their sharpness with time, so fresh knives are prepared each day. A clearance angle of 4° is recommended with glass knives and 12 to 14° with tungsten carbide knives when cutting eyes, although the optimum angle can vary with different microtomes. Fine jewelers’ forceps are used to remove sections from the knife. Sections will adhere to the forceps if their tips are not clean and dry. Sections are mechanically flattened as much as possible before placing them in the water bath. The distilled water in the 28°C water bath must be as clean as possible and is filtered through a 0.22-µm filter to remove dust. Sections are picked up from the water bath with thin tipped brushes and placed on precleaned slides. The slides are briefly dried on a 55°C hot plate and left overnight in an incubator at 45 to 65°C to remove residual moisture. Slides are held in a desiccator until stained.

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SECTION INTERPRETATION Although positioning of the tissue during paraffin or plastic embedding is an initial step in orientation of the eye, careful monitoring during sectioning is necessary for understanding orientation of the final sections. The eye is asymmetrical and the appearance of intraocular structures changes drastically depending on the plane of section and the orientation of the globe. In general, the most useful information is gained by a central sagittal cut passing through the center of the cornea, pupil, lens, and optic nerve. In some situations, sections through several different planes of the eye are needed. To meet both of these objectives, our laboratory routinely cuts modified step sections that we find useful and efficient in revealing the pertinent structural features of the eye. After initial trimming of the tissue block, sections are cut just into the edge of the lens, corresponding to the “A” level of the diagram (Figure 13A.1). Generally, three to four slides with a dozen sections are collected at each level. The diagram also suggests the problems that can occur if orientation is poor and oblique sections are made. Particularly when the thickness of structures is being assessed, oblique sections may suggest that tissue is thicker than it really is (Figures 13A.1 and 13A.2). The appearance of “A, B, and C” sections is illustrated in Figure 13A.2. Even within a specific level, the series of sections must be critically examined. This is particularly true with the optic nerve, since a peripheral section may suggest something quite different from a section through the center of the nerve (Figure 13A.2).

COLLECTION OF OPTIC NERVE Necropsy techniques, including optic nerve removal, were discussed in the previous chapter. We get the best results using the following procedure when studying optic nerve damage in glaucoma. When processing optic nerves, it is important to carefully integrate dissection, fixation, and processing. Immediately after euthanasia, the head is separated from the body. Eyes are removed with gentle dissection, avoiding pulling or pressure on the globe. A triangular piece of skull, having for corners the center of the frontal bone and the two occipital condyles is removed using scissors and forceps. The brain and cerebellum are excised with a small scalpel, leaving a thin layer of brain tissue to protect the optic nerves and to maintain their orientation during fixation. Eyes and heads are fixed in a phosphate-buffered glutaraldehyde paraformaldehyde fixative,3 as described above for plastic processing. The heads are then stored in 0.1 M phosphate buffer pH 7.4 at 4°C. To excise the optic nerves, the remaining brain is removed taking care to avoid stretching of the nerves that lie beneath. A cut is made at the center of the chiasm to separate the nerves. Each nerve is then cut at the optic nerve foramen using fine scissors or scalpel and is carefully removed with microdissection forceps. The nerves are stored in 0.1 M phosphate buffer at 4°C until processed.

EMBEDDING AND SECTIONING OPTIC NERVE To assess axon number and integrity in cross section, optic nerves are placed in a 2% phosphatebuffered paraformaldehyde/glutaraldehyde mixture at 4°C for 2 h and then placed in 2% osmium tetroxide in pH 7.4 phosphate buffer (0.1 M) at 4°C overnight, then rinsed in 0.1 M sodium acetate buffer (for 10 min × 3). The whole nerve is stained in 2% uranyl acetate in 0.1 M sodium acetate buffer for 1 h at 4°C and rinsed in acetate buffer (for 10 min × 3). Tissues are dehydrated in graded ethanol, transferred to propylene oxide, then to a mixture of propylene oxide and resin and embedded in a Polybed 812/Araldite resin. Sections 1 µm thick are cut with a diamond knife and dried on a hot plate at a temperature no lower than 80°C. Sections are stained 28 min in 1% paraphenylenediamine in isopropanol:methanol (1:1),8,9 rinsed in isopropanol:methanol, and mounted with Permount mounting media.*

* Modified protocol, courtesy of Drs. Abbot F. Clark and David Cantu-Crouch, Alcon Laboratories.

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FIGURE 13A.1 Section orientation. A. Step-sectioning levels are indicated by the dotted lines and letters A-E. The solid diagonal line would produce an oblique section, similar to that seen in the accompanying illustration. B. In this oblique section, the pupil (arrow) and central lens are present, but the optic nerve was missed, because of the obliquity. (From Smith, R.S. et al., in Pathology of Genetically Engineered Mice, Word, J. et al., Eds., Iowa State University Press, Ames, 2000, 217. With permission.)

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FIGURE 13A.2 Section and optic nerve orientation. A. In this oblique section, both cornea (C) and retina (R) appear abnormally thickened. The small size of the lens (L) and the appearance of the ciliary body show that the plane of section is peripheral. B. This represents an “A” level section, showing iris and lens, but absence of the pupil and optic nerve. Artifactual wrinkles are present in the retina. C. The lens is of nearly full diameter, but pupil and optic nerve are absent, indicating a “B” level section. D. This “C” level section demonstrates central cornea, pupil, central lens, and optic nerve. E. The optic nerve of a mouse with advanced glaucoma appears relatively normal. F. A deeper cut of the same eye shown in E reveals deep cupping (arrow) of the optic nerve (ON), diagnostic of advanced glaucoma. (Figures 13A.1 and 13A.2 reproduced courtesy of Iowa State University Press from Smith, R.S. et al., Interpretation of ocular pathology in genetically-engineered and spontaneous mutant mice, in Ward, J. et al., Eds., Pathology of Genetically Engineered Mice, University of Iowa Press, Ames, 2000, 217.)

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REFERENCES 1. Luna, L.G., Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology, McGraw-Hill, New York, 1960. 2. Relyea, M.J. et al., Necropsy methods for laboratory mice: biological characterization of a new mutation, in Sundberg, J.P. and Boggess, D., Eds., Systematic Approach to Evaluation of Mouse Mutations, CRC Press, Boca Raton, FL, 1999, 57. 3. Smith, R.S. and Rudt, L.A., Ultrastructural studies of the blood-aqueous barrier, Am. J. Ophthalmol., 76:937, 1973. 4. Schlamp, C.L. and Nickells, R.W., Light and dark causes a shift in the spatial expression of a neuropeptide-processing enzyme in the rat retina, J. Neurosci., 16:2164, 1996. 5. Schlamp, C.L. and Williams, D.S., Myosin V in the retina: localization in the rod photoreceptor synapse, Exp. Eye Res., 63:613, 1996. 6. Hogan, B., Manipulating the Mouse Embryo: A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, 1994. 7. Hoover, S.B. et al., In situ hybridization techniques for studying gene expression in the genetically altered mouse, in Pathology of Genetically Engineered Mice,Ward, J.M. et al., Eds., Iowa State University Press, Ames, 2000, 13. 8. Sadun, A.A., Smith, L., and Kenyon, K.R., Paraphenylenediamine: a new method for tracing human visual pathways, J. Neuropathol. Exp. Neurol., 42:200, 1983. 9. Sadun, A.A. and Schaecter, J.D., Tracing axons in the human brain: a method utilizing light and TEM techniques, J. Electron Microsc. Tech., 2:175, 1985.

B. ELECTRON MICROSCOPY Lesley S. Bechtold and Richard S. Smith Transmission electron microscopy (TEM) is necessary for imaging fine cellular details in mouse eyes. Although obtaining images of the eye is relatively easy, proper preparation of the tissue to ensure that the images are free of artifacts is somewhat more difficult. Portions of the eye, such as the ciliary body, are sensitive to changes in osmolarity and pH, so fixatives and buffers must be chosen carefully to avoid these problems.1 Proper trimming of the tissue is necessary for correct orientation during embedding. Selection of an appropriate resin and attention to details of infiltration is critical in obtaining the highest quality ultrathin sections. This section summarizes techniques that have proved useful for preparing mouse eye tissue for TEM.2

FIXATIVES AND BUFFERS The most commonly used primary fixative for TEM is glutaraldehyde. Glutaraldehyde belongs to a class of fixatives known as noncoagulative or additive fixatives. The purpose of fixation is to preserve the morphology and ultrastructure of cells so that they are as close to the natural state as possible. Glutaraldehyde cross-links but does not coagulate or denature proteins in cells, making protein extraction less likely during dehydration. In addition, gluataraldehyde binds to cellular proteins, adding structural stability. Other noncoagulant fixatives include paraformaldehyde, acrolein, and osmium tetroxide. All act in a similar manner, with some variation in penetration and final tissue contrast. Glutaraldehyde penetrates tissue more slowly than paraformaldehyde, and osmium tetroxide is slower still. For this reason, osmium is usually not used as a primary fixative.3,4 Glutaraldehyde is often employed in conjunction with another fixative. Glutaraldehyde and paraformaldehyde mixtures in phosphate buffer are known as “Karnovsky’s Fix.”5 A useful fixative mixture1 for eyes is 0.8% glutaraldehyde and 1.2% paraformaldehyde in 0.1 M Sorenson’s phosphate buffer at pH 7.2. Higher concentrations of fixatives increase the osmolarity of the final solution. Since some cell membranes are extremely sensitive to high osmolarity, lower concentrations eliminate swelling in tissues such as the

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ciliary body. Freshly opened glass ampoules of concentrated fixatives especially distilled for electron microscopy should be used when preparing fixative mixtures. All fixatives are potential biohazards and should be opened in a well-ventilated hood. Concentrated fixatives are diluted in freshly prepared phosphate buffer made in ultrapure distilled water from a Millipore-type distillation system. All glassware used should be washed and rinsed with the same distilled water. Although several buffers are available, phosphate buffer is the buffer of choice because of its lower osmolarity, compared to cacodylate buffers. The 0.2 M monobasic and dibasic phosphate solutions are diluted with distilled water and concentrated fixative to achieve a final concentration of 0.1 M and a pH of 7.2. The same buffer is used to rinse the tissue after fixation is completed. A second postfixation step with a heavy metal fixative is normally used after primary fixation and trimming of the tissue blocks (see below). Osmium tetroxide, a yellow crystal that dissolves slowly in aqueous systems, is generally used to postfix tissues. The electron-dense osmium provides added contrast for resolution of organelles and membrane structures in the eye. Osmium also acts as a mordant for lead stains that improves contrast. Thorough washing with buffer between fixatives is necessary to avoid reactions with aldehyde fixatives that can cause electron-dense tissue precipitates. All fixation and rinsing steps are performed at 4°C. Although a cooler temperature slows the rate at which fixative enters the tissue, it also minimizes shrinkage artifacts and extraction.

FIXATION AND TISSUE TRIMMING Two methods are commonly used for tissue fixation for electron microscopy. Vascular perfusion by the intracardiac route is performed using a clearing solution, such as phosphate-buffered saline (PBS), to remove blood from the circulatory system. The PBS solution is then replaced with freshly prepared fixative and between 4 and 8 ml of fixative are perfused. This technique is often useful for fixation of brain and other large organs. However, comparative study of immersion and perfusionfixed eye tissue suggests that tissue morphology is equivalent with both methods. The preferred method for fixing eyes is immersion. The tissue is placed in fresh fixative at 4°C as soon as the eye is removed. The concept of making a small opening in the eye prior to fixation has been suggested to allow better penetration of fixative. However, this may induce partial collapse of the globe and that could damage structures within the eye. The mouse sclera is so thin that fixative penetration is usually not a problem. The timing of eye dissection after initial fixation depends on which ocular tissue is being collected. Anterior segment structures are relatively resistant to damage during dissection. Satisfactory results are obtained if the undisturbed whole eye is fixed for 60 to 90 min. The retina requires a longer preliminary fixation time because it tends to detach spontaneously from the retinal pigment epithelium, eliminating normal anatomic relationships. Before opening the eye to cut retinal tissue blocks, the retina should be fixed for at least 2 to 3 h. Fixation times as long as 12 h still give excellent anatomic preservation of the outer retina and retinal pigment epithelium, although there is deterioration of membrane contrast in the inner retina. For the best overall preservation of all retinal layers, the eye should be opened after 3 h of preliminary fixation. A fine-pointed disposable blade is used to make an opening posterior to the ciliary body and a small Vannas-style scissors is used to extend the cut around the eye, separating anterior and posterior segments (Figure 13B.1). The exact location of the cut is varied depending on whether anterior or posterior structures will be collected. The lens usually remains attached to the anterior segment and must be gently removed before anterior segment tissue blocks are prepared. Since tissue size has important effects on fixation quality, the ocular tissues must be trimmed to a size small enough for proper postfixation and infiltration to occur. Tissue blocks of anterior segment are made by placing the cornea down on the dissecting surface and dividing the anterior segment into two pieces, using a clean fresh razor blade. If the cornea is being collected, this should be

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done gently to avoid damage to the corneal epithelium. The posterior segment is divided in half in similar fashion with the sclera facing the dissecting surface. In both instances, the hemisected segment is cut into small wedges in the shape of an isosceles triangle that measure about 0.5 × 1.5 mm. This shape is helpful for achieving the desired orientation during embedding, as well as assisting in tissue orientation. For example, if the retina is cut so that the apex of the triangle points toward the optic nerve, more peripheral retina can be identified. Figure 13B.1 shows diagrammatically the steps in tissue trimming for the anterior segment.

FIGURE 13B.1 A. Fine forceps and curved Vannas-style scissors are used to divide the globe (dotted lines, A) into anterior and posterior segments (B and C). The position of this cut determines whether the lens remains with the anterior or posterior segment. After removal of the lens the anterior segment is divided into pie-shaped wedges (dotted white lines, B) to produce small triangles that include cornea, iris, and ciliary body (D).

POSTFIXATION AND EMBEDDING Once the ocular tissues are trimmed, additional fixation is done using the primary aldehyde fixative. The appropriate length of fixation at 4°C can vary from 4 h to overnight. Two or three 15-min washes with cold phosphate buffer are performed before postfixation with osmium tetroxide. A 2% aqueous osmium solution is combined with buffer to make a 1% solution. The tissue is postfixed for 2 to 4 h at 4°C, washed two or three times with buffer, and dehydrated through graded ethanols. The purpose of dehydration and infiltration is to remove all traces of water from the tissue and to replace it with embedding medium. The melanosomes of iris, ciliary body, choroid, and retinal pigment epithelium are particularly difficult to infiltrate and, if infiltration is not optimal, are prone to shatter during sectioning. The type of resin used is the choice of the investigator, but epoxy resins such as Epon, Araldite, or Spurrs produce the best results with ocular tissue and are stable in the electron beam. The components are easily obtained from any electron microscopy supplier and have a long shelf life. For eyes, an Epon–Araldite mixture,6 consistently provides sections with almost no chatter, wrinkles or tears due to the melanosomes. Although the older resin component Epon is no longer available, its substitute, Embed 812 (Electron Microscopy Sciences, Fort Washington, PA), is similar to the original resin. Adequate infiltration of ocular tissue requires up to 48 h and should be carried out on a rotator. As the resin and ethanol are not readily miscible, a transfer solvent, such a propylene oxide, is used to promote infiltration of the tissue with resin.

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FIGURE 13B.2 After fixation, the tissues are embedded in resin in a flexible mold and the resin is polymerized. Tissue is oriented in the plastic block as shown. The end view of the tissue shows its position in the microtome chuck. The face of the block would be trimmed to a truncated pyramidal shape prior to cutting.

Once the tissue is completely infiltrated, it must be properly oriented in small, flat embedding molds (Figure 13B.2). The standard blue-colored, silicon rubber, flat embedding molds from Ladd Research (Burlington, VT) are satisfactory. The mold depressions are filled with a small amount of resin. A tissue wedge is oriented at the tapered end of the depression by placing the wedge curled edges up and the body of the wedge flat on the bottom of the mold depression with a long cut edge up flat against the tapered point of the depression (Figure 13B.2). The molds are placed in an oven at the appropriate temperature for the resin used and for the required amount of time to polymerize the blocks completely. The tapered ends of the blocks are trimmed to produce a trapezoidal face so that a cross section of the tissue is obtained. Blocks are sectioned on an ultramicrotome to produce 80-nm-thick sections. These sections are picked up on uncoated, 300-mesh copper grids. The grids are stained with uranyl acetate and counter-stained with Reynolds lead citrate.7 As with all solutions used to prepare eyes for TEM, stains work best if freshly prepared. Grids are stained for 30 to 60 min with each stain and washed thoroughly with distilled water after each stain. Water is removed from the grid using a piece of filter paper after the final wash and the grids are air-dried before they are examined in the transmission electron microscope.

ROUTINE PROTOCOLS Production of high-quality electron micrographs requires equal portions of science, art, and technical skill. Although there are many variations favored by different investigators, the specific procedures successfully employed for fixation of mouse eyes in our laboratories at The Jackson Laboratory are included below. Fixation The solution used consists of 1.2% paraformaldehyde and 0.8% glutaraldehyde in 0.1 M phosphate buffer, pH 7.2, and fixation is performed overnight at 4ºC. Specimens are washed twice for 15 min with 0.1 M phosphate buffer at 4ºC. Specimens are allowed to warm to room temperature during the second wash. Tissue is postfixed in 1% osmium tetroxide in phosphate buffer for 4 h at 4ºC.

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Dehydration This procedure is performed at room temperature for 15 min in each ethanol concentration: 60, 80, 95%; and twice for 15 min in 100% ethanol. Propylene oxide is used as a transition solvent between absolute alcohol and the plastic resin; the sample is placed in two changes of propylene oxide for 10 min each. Infiltration and Embedding The tissue is placed in a 1:1 mixture of propylene oxide: resin for 18 to 24 h on a rotator at room temperature. This is followed by 24 h of pure resin on a rotator. Fresh resin is used to embed specimens in molds (see Figure 13B.2). The resin is polymerized at 65ºC for 72 h. Solution Preparation 1. Phosphate Buffer, pH 7.2 Solution A—0.2 M sodium phosphate, monobasic: dissolve 27.6 g of NaH2PO4·2H2O in 1000 ml of distilled water. Solution B—0.2 M sodium phosphate, dibasic: dissolve 35.61 g of NaH2PO4·2H2O in 1000 ml of distilled water. Prepare 2000 ml of 0.1 M buffer by combining 280 ml of solution A, 720 ml of solution B, and 1000 ml of distilled water. Check pH and adjust as necessary with 1 N NaOH or 1 M HCl. 2. Epon–Araldite Resin Using a disposable 100-ml plastic tripour beaker, combine the following ingredients on a stir plate for at least 30 min: 14 ml DDSA (dodecenyl succinic anhydride) 6 ml Embed 812 4 ml Araldite 506 1 ml DBP (dibutyl phthalate) 0.7 ml DMP-30 (accelerator) This resin can be mixed with propylene oxide during infiltration and used full strength for embedding. It will polymerize at 65ºC. 3. Uranyl Acetate Stain Dissolve 1 g of uranyl acetate powder in 50 ml of distilled water. Stain grids for up to 1 h with this stain. Rinse well with distilled water after use. 4. Lead Citrate Counterstain Dissolve 1.33 g of lead nitrate and 1.76 g of sodium citrate in 30 ml of distilled water. Mix well on a stir plate for 30 min. Add 10 ml of 1 N NaOH. Mix for an additional 5 min. Add 10 ml of distilled water, mix, and store at 4ºC until used. Stain grids for up to 1 h at room temperature with this stain. Fresh stain will result in less lead precipitate on the section.

REFERENCES 1. Smith, R.S. and Rudt, L.A., Ultrastructural studies of the blood-aqueous barrier 2: the barrier to horseradish peroxidase in primates, Am. J. Ophthalmol., 76: 937, 1973. 2. Bechtold, L.S., Ultrastructural evaluation of mouse mutations in Systematic Approach to Evaluation of Mouse Mutations, Sundberg, J.P. and Boggess, D., Eds., CRC Press, Boca Raton, FL, 2000. 3. Bechtold, L., Why use osmium tetroxide? Microsc. Soc. Canada Bull., 29:16, 2001. 4. Hayat, M.A., Fixation for Electron Microscopy, Academic Press, New York, 1981, 148–179.

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5. Karnovsky, M.J., A formaldehyde-glutaraldehyde fixative of high osmolarity for use in electron microscopy, J. Cell Biol., 27:137A, 1965. 6. Mollenhauer, H.H., Plastic embedding mixtures for use in electron microscopy, Stain Technol., 39:111, 1964. 7. Reynolds, E.S., The use of lead citrate at high pH as an electron opaque stain in electron microscopy, J. Cell Biol., 17:208, 1964.

SUGGESTED READING Glauert, A.M., Fixation, Dehydration and Embedding of Biological Specimens, Elsevier North-Holland Biomedical Press, Amsterdam, the Netherlands, 1975. Hayat, M.A., Principles and Techniques of Electron Microscopy, Biological Applications, 3rd ed., CRC Press, Boca Raton, FL, 1989.

C. IMMUNOHISTOCHEMISTRY Sakae Ikeda, Melissa J. Relyea, and John P. Sundberg

INTRODUCTION Immunohistochemistry, a method for determining localization of proteins within a tissue, is an important tool in eye research. The method is used to determine where in the eye a protein is expressed (i.e., retina), which cell types within the structure contain the protein (i.e., photoreceptor cells), and its location within a specific cell (i.e., outer segments). Immunohistochemical localization is frequently useful in elucidating protein function within the eye. Furthermore, improper expression of proteins may identify specific abnormalities as a result of a mutation. The basic principles of immunohistochemistry are quite simple. An antibody that specifically binds to a protein on the tissue section is localized by using a detection reagent labeled with a fluorescent dye or an enzyme that reacts with a chromogen. Since many variables in the procedure (such as choice of primary antibodies, dilution of antibodies, fixation methods of the tissue, etc.) can easily alter the final results, the methodology is not as simple as it may appear. In this chapter, major factors that affect immunohistochemical studies are discussed and some general protocols for staining eye sections are provided. More detailed information on methodology and applications of immunohistochemistry are provided elsewhere.1-6

PRIMARY ANTIBODIES AND FIXATION METHODS The two most important factors for success of an immunohistochemical procedure are the choice of primary antibody and the method chosen for tissue fixation. Although the goal is to achieve specific antibody labeling while preserving the morphology of the tissue, strong fixation often disrupts the antigenic sites of a protein, which leads to a weak or absent signal. A balance between these two factors is necessary and must be evaluated for each tissue and each antibody. It is important to choose primary antibodies that specifically bind to the protein of interest in the species being studied. For example, an antibody raised against a human protein may not react with the corresponding mouse protein. As an initial step, the antibody should be tested on a Western blot of eye lysate for specificity. Observation of a major band of expected size with no background is a good indication that the antibody will be useful, since major cross-reactivity is detected by this method. The next step in testing antibody specificity is to include a negative control for the procedure. The primary antibody should be replaced, ideally by preimmune serum from the same animal in which the antibody was raised or by normal serum from an animal of the same species. This control distinguishes a real signal from background staining produced by nonspecific binding of secondary reagents. Another approach for detecting nonspecific background staining is to preadsorb the

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antibody with excess amounts of antigen against which the antibody was raised prior to applying it on the tissue section. An absence of signal after preadsorption indicates that the signal is specific to the antigen.7 Different fixation and embedding methods should be considered depending on antibody characteristics. The mildest preservation method is to use unfixed or briefly fixed frozen tissue. The advantage of this procedure is that the epitope (the antigenic determinant on an antigen to which an antibody binds) is more likely to be preserved; the disadvantage is that tissue morphology is poorly preserved. In contrast, fixed paraffin-embedded tissues produce superior morphology. The disadvantage of using standard fixation and paraffin-embedding techniques is that they may disrupt or mask epitopes. For example, tissues fixed in the commonly used 10% neutral buffered formalin solution often remain in the fixative for long periods of time. As long as tissues are in the fixative, amino groups will be cross-linked, thereby changing the tertiary structure of the epitopes. Simply transferring tissues after overnight fixation into 70% ethanol may be enough to maintain epitope integrity. For some antibodies, antigen retrieval methods discussed later may be useful in dealing with this problem. The availability of effective antibodies for a specific protein of interest should be evaluated by a thorough literature search and by reviewing available data from commercial sources. Antibodies that work in some applications, such as Western blotting, may or may not work well in immunohistochemistry. It is important to know the applications for which an antibody works, what methods for tissue preparation have been tested (frozen or paraffin), and whether dilution of the antibody is recommended for immunohistochemistry. Although manufacturer’s data may not always be complete or correct, it provides a starting point for developing new techniques. If there is little information about an antibody, it should be tested on both frozen and paraffin sections and with a variety of fixatives to achieve the best conditions for high-quality results.

ENZYME AND FLUORESCENCE DETECTION Both enzyme and fluorescence methods are commonly used to detect antibody binding. Although good results are achieved with either method, each has its own advantages that make it suitable for particular applications. Enzyme Detection This method, when combined with the avidin–biotin system, has high sensitivity.8,9 Following the application of a biotinylated secondary antibody, avidin/biotinylated enzyme complex (ABC) is applied to amplify the signal. The bound enzyme complex is next incubated with a substrate to produce a colored reaction product that localizes the signal. The tissue is usually counterstained with a standard nuclear staining reagent such as hematoxylin and can be viewed by light microscopy. The signal and the tissue morphology can be viewed simultaneously, which makes it easier to determine where the protein is expressed in the tissue. Figure 13C.1 shows mouse retina stained with an antibody against tubby-like protein 1 using a peroxidase detection system and diaminobenzidine (DAB) as the substrate. Brown staining is observed primarily in the outer plexiform layer (OPL) and inner segments (IS) of the retina.10 Another advantage of this method is that a substrate capable of producing an insoluble reaction product, such as DAB, can be chosen. This enables permanent mounting of the slide with organic-solvent-based mounting medium, which is an important feature to consider if permanent slide storage is desired.

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FIGURE 13C.1 Normal mouse retina. The retina is stained with an antibody directed against tubby-like protein 1, using a peroxidase detection system with DAB as the substrate. The outer plexiform (OPL) and photoreceptor inner segments (IS) demonstrate a brown reaction product. The apparent tear in the RPE (*) is an artifact.

FIGURE 13C.2 Normal mouse retina. The retina is stained with an anti-rhodopsin antibody that was detected using a biotinylated secondary antibody conjugated with FITC. FITC and DAPI signals were viewed separately and the images merged. The outer segments (OS) are a brilliant green, whereas the inner nuclear (INL), outer nuclear layers (ONL), and the inner segments (IS) are not labeled.

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Fluorescence Detection Immunofluorescence employs a secondary antibody or avidin labeled with a fluorochrome for signal detection. Widely used fluorochromes include fluorescein isothiocyanate (FITC, green), tetramethyl rhodamine (TRITC, red), and their derivatives. The tissue section is viewed under a fluorescence microscope using an appropriate filter for each fluorochrome. Figure 13C.2 is a paraffin section of retina stained with an antibody against rhodopsin in which the signal was detected using a biotinylated secondary antibody and avidin conjugated with FITC. FITC and DAPI (4,6-diamidino-2phenylindole) signals were viewed separately using appropriate filters and the collected images were merged. This method gives better signal resolution and is especially suitable for viewing at higher magnification and for reconstructing three-dimensional structure, using confocal microscopy. Immunofluorescence is also useful for simultaneous localization of two or more antigens on a single tissue section. Primary antibodies raised in different species are detected with reagents labeled with different fluorochromes. Each signal is viewed sequentially using an appropriate filter and the collected images are combined to show colocalization or different patterns of localization for multiple proteins.11 Because fluorochrome-labeled reagents are incompatible with permanent mounting media, longtime storage of slides is unsatisfactory. In addition, the fluorochromes fade under light in time. Recently developed nonbleaching fluorochromes offer alternatives that avoid rapid loss of signal. Examples of nonbleaching reagents include indocarbocyanine Cy-2 (FITC substitute) and indocarbocyanine Cy-3 (rhodamine substitute), from Jackson ImmunoResearch (West Grove, PA) or Chemicon (Temecula, CA), and Alexa 488 (FITC substitute) and Alexa 568 (rhodamine substitute), from Molecular Probes (Eugene, OR). It has been suggested that formalin-fixed tissues are prone to nonspecific fluorescence and because of that, nonfixed frozen tissues should be used for immunofluorescence. However, we have not encountered major problems using paraformaldehyde-fixed paraffin embedded tissues for fluorescence detection systems.12-14 This is likely due to the use of modern interference filters that are more selective in terms of excitation and emission wavelengths than those on earlier fluorescence microscopes. Use of more selective filters minimises detection of non-specific background signals.15

OVERVIEW OF THE STAINING PROCEDURE Tissue Preparation For frozen sections, the eye is embedded in OCT compound (Miles Laboratories, Elkhart, IN). Prior to embedding, the eye can be immersion-fixed in 4% paraformaldehyde (PFA) for a short time to achieve better morphology. Frozen sections are cut on a cryostat and dried at room temperature for 1 to 12 h and then fixed in acetone or PFA before processing. After drying, the sections are stored at –20°C or below (preferably –80°C), wrapped separately in aluminum foil or plastic, and sealed in an airtight container with desiccant. Before use, the sections should be warmed to room temperature in the container to prevent condensation.16 For producing paraffin sections, the eye is immersion fixed in 4% PFA in phosphate-buffered saline (PBS) or in alternative fixatives such as Bouin’s or Telly’s and processed for routine paraffin embedding (see Section 13A). It is often useful to test different fixatives, because some antigens are better preserved with a specific fixative.16 To prepare for immunohistochemical staining, sections are deparaffinized in xylene and rehydrated through graded alcohols to water. As previously mentioned, fixation and paraffin embedding often produce severe masking of antigenic sites, likely due to protein cross-linking. Several methods have been developed to overcome this problem.17 The most commonly used technique is heating, using either microwave ovens or pressure cookers. Although this technique has been successfully used with many antibodies, optimal

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conditions (especially the buffer pH and the temperature) vary between antibodies. Accordingly, different conditions should be tested to achieve the optimal results for each antibody. Nonspecific binding of antibodies can be blocked by incubation of the tissue with normal serum from the animal in which the secondary antibody was generated, prior to applying the primary antibody to tissue sections. Ovalbumin or bovine serum albumin (BSA) are also useful for blocking nonspecific reactions. With peroxidase-labeled detection techniques, endogenous tissue peroxidase activity can be minimized by incubating slides in 3% H2O2 for 15 to 30 min prior to antibody incubation. Incubation with Antibodies The immunohistochemical reaction begins with incubation of tissue sections with a primary antibody directed against the antigen of interest. Titering (determining appropriate antibody dilution) of the primary antibody is very important to achieve an intense signal without producing nonspecific reactions. When used for the first time, the antibody should be tested at several different dilutions, although manufacturers’ recommendations are a useful starting point. Sections are incubated with the primary antibody solution containing a blocking reagent such as normal serum or BSA to avoid nonspecific binding. Incubation is carried out in a humidified chamber. We routinely perform 2-h incubations at room temperature or overnight incubation at 4°C. The sections are next incubated with a labeled secondary reagent. The label chosen should be compatible with the detection system. For enzyme detection, the avidin–biotin system8,9 that allows for amplification of signal is commonly employed. Kits are available from most vendors. A biotin-labeled secondary antibody against the IgG of the species in which the primary antibody was generated is used. For immunofluorescence, a fluorochrome-labeled secondary antibody is used. If amplification of signal is desired using an avidin-biotin system, a biotinylated secondary antibody can be utilized, followed by detection with avidin conjugated with a fluorochrome. The appropriate dilution of the secondary antibody should also be determined to achieve optimal staining without background. Enzyme Detection Following application of the biotinylated secondary antibody, ABC is applied to amplify the signal. The commonly used enzyme systems are either horseradish peroxidase or alkaline phosphatase. Peroxidase-based systems are most commonly used because of their sensitivity, dense reaction product, and stability. Alkaline phosphatase systems are useful if endogenous peroxidase activity interferes with data interpretation. However, alkaline phosphatase results are usually less intense than those obtained with peroxidase.16 The signal is visualized by adding an enzyme substrate. The most commonly used peroxidase substrates are DAB and 3-amino-9-ethylcarbazole (AEC). DAB produces a brown precipitate that is insoluble in organic solvents, contrasts well with many counterstains, and can be used with organic solvent–based mounting media. In pigmented tissue, care should be taken not to confuse DAB precipitate with melanin pigment. AEC produces a bright red stain that contrasts well with tissue components when hematoxylin is used as a counterstain. AEC is alcohol soluble, so aqueous-based mounting media must be used. Alkaline phosphatase substrates include nitro blue tetrazolium (BCIP/NBT), iodoblue tetrazolium (BCIP/INT), and Fast Red/Napthol AS-Mx. BCIP/NBT and BCIP/INT are both compatible with organic solvents, whereas Fast Red is alcohol soluble.16 When the enzyme reaction is complete, the tissue can be counterstained to help identify cells and tissue structure and to provide contrast to the reaction product utilized in the immunohistochemical reaction. Commonly used counterstains include hematoxylin (blue), toluidine blue (blue), light green (green), nuclear fast red (red), and methyl blue (blue). The choice of counterstain is a subjective one, but should be compatible with both the substrate color and the mounting medium.16

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Fluorescence Detection After application of a fluorochrome-labeled secondary antibody or avidin conjugated with a fluorochrome, tissue sections can be mounted with an anti-photobleaching mounting medium such as Vectashield Mounting Medium (Vector Laboratories, Burlingame, CA) or SlowFade Light Antifade solution (Molecular Probes, Eugene, OR), which help prevent rapid loss of fluorescence. If staining of the cell nuclei is desired, DNA-specific fluorescence dyes such as 4,6-diamidino-2-phenylindole (DAPI, blue) or proiodide (PI, red) can be applied prior to mounting. Although the slides can be stored in the dark at 4°C, the signal should be viewed under a fluorescence microscope as soon as possible. Since permanent mounting of the slides is impossible, it is important to collect images immediately.

GENERAL PROTOCOLS FOR IMMUNOHISTOCHEMISTRY Although there are many specific techniques published, we have found the following to be particularly useful. Circumstances alter cases, and these are intended only as guidelines that the reader may find useful. Preparation of paraffin sections: 1. Deparaffinize and rehydrate sections through graded series of alcohol to water. For peroxidase detection, incubate sections with 3% hydrogen peroxide in methanol for 15 min after incubation with absolute ethanol during the rehydration process. 2. Wash in PBS for 5 min. 3. Incubate sections with diluted normal serum for 20 min. 4. Blot excess serum from sections. Sections are ready to be processed. Preparation of frozen sections: 1. Air-dry the sections. 2. Fix in acetone for 10 min. 3. Wash in PBS for 5 min. 4. For peroxidase detection, incubate sections with 3% hydrogen peroxide in water for 15 min. 5. Wash in PBS for 5 min. 6. Incubate sections with diluted normal serum for 20 min. 7. Blot excess serum from sections. Sections are ready to be processed. Immunohistochemistry using enzyme detection: 1. Incubate sections with the primary antibody diluted in PBS with normal serum for 2 h at room temperature or overnight at 4°C in a humidified chamber. 2. Wash in PBS for 5 min. 3. Incubate sections with the biotinylated secondary antibody diluted in PBS with normal serum for 30 min at room temperature in a humidified chamber. 4. Wash in PBS for 5 min. 5. Incubate sections with ABC Reagent for 45 min at room temperature in a humidified chamber. 6. Wash in PBS for 5 min. 7. Incubate sections in the substrate solution until proper color development occurs. 8. Rinse sections in tap water. 9. Counterstain and mount sections. Immunofluorescence: 1. Incubate sections with the primary antibody diluted in PBS with normal serum for 2 h at room temperature or overnight at 4°C in a humidified chamber.

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2. Wash in PBS for 5 min. 3. Incubate sections with a fluorochrome conjugated or biotinylated secondary antibody diluted in PBS with normal serum for 30 min at room temperature in a humidified chamber. 4. Wash in PBS for 5 min. 5. If a biotinylated secondary antibody is used, incubate sections with fluorochrome conjugated avidin diluted in PBS. 6. Wash in PBS for 5 min. 7. Stain nuclei with DAPI. 8. Wash in PBS for 5 min. 9. Mount sections with anti-photobleaching mounting media.

INTERPRETATION OF RESULTS Once the reaction is completed, it is important to determine if this is indeed a signal achieved by specific binding of the primary antibody. Apparent signals may not indicate antigen expression or localization, but may instead indicate nonspecific antibody binding or even cross-reaction to an epitope of another antigen. To distinguish a real signal from a false one, proper controls should be included in an experiment. Positive Controls To confirm that the procedure was performed properly and that the reagents were working correctly, sections of tissue known to express the protein at relatively high levels should always be included as controls. The positive control tissue should be fixed and processed in exactly the same manner as the test samples, because this can indicate if the fixation and embedding methods were compatible with the primary antibody used. Negative Controls It is also important to include negative controls to confirm the specificity of the reagents used in the experiment. As mentioned earlier, a section incubated with the primary antibody preadsorbed with excess amount of the purified antigen is a valuable negative control to show specificity of the antibody. When a polyclonal antibody is used, another important control is to use serum from the same animal in which the antibody was generated, although nonimmmune serum from an animal of the same species is usually acceptable, especially when using inbred mice raised and maintained in pathogen-free conditions. A section of tissue that does not express the antigen is also a good negative control. In this regard, tissue from a mutant mouse lacking the protein under study, if available, would serve as the best negative control.12 In addition, a negative control slide incubated only with the secondary reagents should be included. Results from this control will indicate if any of the staining is due to nonspecific binding of the secondary reagents. Autofluorescence The interpretation of immunofluorescence can be complicated by the presence of autofluorescent pigments in the tissue, such as lipofuscin in the retinal pigment epithelium (RPE). Lipofuscin accumulates in the RPE with age in mammalian eyes, and fluoresces intensely at the spectrum overlapping those of commonly used fluorophores. However, in most cases, this problem can be overcome by including a control slide without the fluorescent-labeled reagents. By comparing the staining pattern with this control, it is fairly easy to determine which signal is specific and not caused by autofluorescence. If the protein of interest is expressed where strong autofluorescence is observed,

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switching to an enzyme detection system where autofluorescence is not a problem should be considered. A second approach is to treat the tissue sections with chemicals such as cupric sulfate or Sudan Black B that can reduce or eliminate lipofuscin autofluorescence, while retaining the specific immunohistochemical signal.18 Endogenous Enzyme Activity Some cells have high levels of endogenous peroxidase and will give false positive reactions with routine peroxidase enzyme detection systems. Endogenous peroxidase activity may be blocked by incubation of slides in a solution of hydrogen peroxide. If nonspecific staining due to endogenous peroxidase continues to cause difficulty, changing the enzyme system used should eliminate the problem.

REFERENCES 1. Polak, J.M. and Van Noorden, S., Introduction to Immunocytochemistry, 2nd ed., Springer-Verlag, New York, 1997. 2. Bullock, G.R. and Petrusz, P., Techniques in Immunocytochemistry, Vol. 1, Academic Press, London, 1982. 3. Bullock, G.R. and Petrusz, P., Techniques in Immunocytochemistry, Vol. 2, Academic Press, London, 1983. 4. Bullock, G.R. and Petrusz, P., Techniques in Immunocytochemistry, Vol. 3, Academic Press, London, 1986. 5. Bullock, G.R. and Petrusz, P., Techniques in Immunocytochemistry, Vol. 4, Academic Press, London, 1989. 6. Sternberger, L.A., Immunocytochemistry, 3rd ed., John Wiley & Sons, New York, 1986. 7. He, W. et al., GFP-tagged expression and immunohistochemical studies to determine the subcellular localization of the tubby gene family members, Brain Res. Mol. Brain Res., 81:109, 2000. 8. Guesdon, J.L., Ternynck, T., and Avrameas, S., The use of avidin-biotin interaction in immunoenzymatic techniques, J. Histochem. Cytochem., 27:1131, 1979. 9. Hsu, S.M., Raine, L., and Fanger, H., Use of avidin-biotin-peroxidase complex (ABC) in immunoperoxidase techniques: a comparison between ABC and unlabeled antibody (PAP) procedures, J. Histochem. Cytochem., 29:577, 1981. 10. Ikeda, S. et al., Cell-specific expression of tubby gene family members (tub, Tulp1,2, and 3) in the retina, Invest. Ophthalmol. Vis. Sci., 40:2706, 1999. 11. Applebury, M.L. et al., The murine cone photoreceptor: a single cone type expresses both S and M opsins with retinal spatial patterning, Neuron, 27:513, 2000. 12. Ikeda, S. et al., Retinal degeneration but not obesity is observed in null mutants of the tubby-like protein 1 gene, Hum. Mol. Genet., 9:155, 2000. 13. Ikeda, A. et al., Neural tube defects and neuroepithelial cell death in Tulp3 knockout mice, Hum. Mol. Genet., 10:1325, 2001. 14. Haider, N.B., Naggert, J.K., and Nishina, P.M., Excess cone cell proliferation due to lack of a functional NR2E3 causes retinal dysplasia and degeneration in rd7/rd7 mice, Hum. Mol. Genet., 10:1619, 2001. 15. Mason, D.Y., Micklem, K., and Jones, M., Double immunofluorescence labelling of routinely processed paraffin sections, J. Pathol., 191:452, 2000. 16. Relyea, M.J., Sundberg, J.P., and Ward, J.M., Immunohistochemical and immunofluorescence methods, in Systematic Approach to Evaluation of Mouse Mutations, Sundberg, J.P. and Boggess, D., Eds., CRC Press, Boca Raton, FL, 1999, 131. 17. Shi, S.R., Cote, R.J., and Taylor, C.R. Antigen retrieval immunohistochemistry: past, present, and future, J. Histochem. Cytochem., 45:327, 1997. 18. Schnell, S.A., Staines, W.A., and Wessendorf, M.W., Reduction of lipofuscin-like autofluorescence in fluorescently labeled tissue, J. Histochem. Cytochem., 47:719, 1999.

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D. NORTHERN AND IN SITU HYBRIDIZATION Sakae Ikeda, Chia-Yang Liu, and Winston W.-Y. Kao

INTRODUCTION Northern and in situ hybridization are common techniques used to detect levels of expression of specific mRNAs and to identify cell types that express a gene of interest. Northern hybridization can provide semiquantitative information about the levels of specific mRNA expression in tissues at various stages of embryonic development. This is achieved by cohybridization of the blot with 32 P-labeled cDNA of a generally expressed gene, such as Gapd (glyceraldehyde-3-phosphate dehydrogenase), and comparing the intensity of signals.1 As an example of the applications of these two techniques, Northern hybridization has shown that the lumican gene is ubiquitously expressed in many organ systems, such as skin, heart, eye, and muscle. The levels of lumican expression are higher during embryonic development than in adults. Northern hybridization cannot identify cell types in which the gene is expressed when the source tissue consists of multiple cell types, such as eye, skin, and heart. Cellular localization can be achieved by in situ hybridization. For example, it has been demonstrated that the lumican gene is expressed by mesenchymal cells in skin, heart, eye, and lung of normal adult animals, the neural retina during development, and the corneal epithelial cells during wound healing.1,2 In situ hybridization can also be used to label cells of specific cell lineages to follow their migration during embryonic development. For example, in situ hybridization using keratocan riboprobes indicates that periocular mesenchyme of neural crest origin may give rise to corneal keratocytes, iris fibroblasts, eyelid stroma, and trabecular meshwork.3 Thus, the combination of Northern and in situ hybridization can produce useful information regarding levels and spatial-temporal expression patterns of mRNAs during embryonic development and in adults. The procedure of Northern hybridization is straightforward and most reagents are commercially available. The techniques of in situ hybridization are more labor intensive and require tissue sectioning. It should be noted that there are other techniques that permit more thorough analysis of gene expression patterns, such as DNA microarray studies,4 accurate determination of mRNA levels (by real-time RT-PCR, or reverse transcription polymerase chain reaction),5,6 and RNase protection assays.7 This chapter focuses on the techniques and procedures for Northern blotting and in situ hybridization.

PRECAUTIONS AGAINST RIBONUCLEASES In both Northern and in situ hybridization procedures, it is critical to avoid degradation of RNA by ribonucleases (RNases) that are stable enzymes present in tissues. Every effort should be made to assure that all glassware used in experiments is RNase-free. This can be accomplished by baking or treating glassware with alkali (2 M NaOH). RNase-free solutions are prepared by treating with 0.1% diethylpyrocarbonate (DEPC) with agitation overnight followed by autoclaving for 40 min. Solutions to be buffered with Tris should be prepared with DEPC-treated water prior to adding the Tris, as Tris inactivates DEPC. Gloves should be worn and aerosol barrier tips used for pipetting to avoid possible contamination with RNases from external sources.

NORTHERN HYBRIDIZATION Northern hybridization is a method for detecting specific sequences in RNA preparations blotted onto a solid support. RNA that is fractionated by gel electrophoresis is transferred to a membrane and subjected to hybridization with labeled probes.

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Total RNA Preparation Protocols for preparing total RNA from tissue involve cell lysis in the presence of a strong protein denaturant that inhibits RNase activity. The most widely used denaturing procedure is the single-step method using guanidine thiocyanate.8 This technique is based on the ability of RNA to remain in the water-soluble phase that contains guanidine thiocyanate under acidic conditions, while most proteins are found in the phenol/chloroform organic phase. Large fragments of DNA remain in the interface under these conditions. This method is rapid, can be used to prepare several samples at the same time, and is suitable for isolation of RNA from small quantities of tissue. Many RNA preparation kits that utilize this method are commercially available, such as Tri-Reagent (Molecular Research Center, Inc., Cincinnati, OH),9 and produce satisfactory results. If mRNA of low abundance is to be detected, poly (A)+ RNA should be isolated from total RNA and used for Northern analysis. Because most mRNAs contain a poly (A) tail, this fraction is enriched for mRNA. A general protocol can be found elsewhere10 and commercial kits are also available. Protocols for Northern Hybridization Preparation of Total RNA Frozen tissue (at –70°C) is homogenized in ice-cold Tri-Reagent at 20 ml tissue (or 1 ml mouse eye) with a high-speed homogenizer system (Cole–Palmer Instrument Co., Vernon Hills, IL). Tissue debris is removed by centrifugation at 8000 g for 10 min. To the supernatant, one-fifth volume of chloroform (0.2 ml/ml Tri-Reagent) is added. The mixture is vigorously mixed with a Vortex. After centrifugation at 8000 g for 10 min, RNA is recovered from the aqueous phase by precipitation with an equal volume of isopropanol added to the solution followed by incubation at –20°C for at least 60 min. Samples are centrifuged at 10 to 14,000 rpm for 10 min in a microcentrifuge. The isopropanol is decanted and the RNA pellet is air-dried. The RNA is then dissolved in DEPC-treated water and stored at –70°C until use. For long-term storage, RNA can be ethanol precipitated and kept at –70°C. Agarose Gel Electroporesis Isolated total or poly(A)+ RNA is fractionated on agarose gel and transferred to a membrane support (Northern blotting). The blot is hybridized with labeled DNA or RNA probes to detect specific sequences. Since RNA is single-stranded and prone to form secondary structures, RNA samples must be electrophoresed under denaturing conditions to obtain good resolution. A denaturant that is commonly used for RNA is formaldehyde,11 which is added to the gel and the sample during gel electrophoresis. After electrophoresis, the gel should be stained with ethidium bromide, and examined, using an ultraviolet transilluminator. Ribosomal RNA (rRNA) molecules are visualized as sharp bands (28S: 4718bp; 18S: 1874bp) if the RNA samples are of good quality. These rRNA bands can also be used as internal size markers. When poly(A)+ RNA is fractionated, RNA molecular weight markers should be used to establish transcript sizes. Preparation of 1% agarose gel: 1. Melt 1 g agarose in 73 ml water plus 10 ml 10× MOPS buffer. 2. Cool the solution to 65°C; add 17 ml of 37% formaldehyde. 3. Pour the gel solution onto a tray with appropriate sample well combs. Sample preparation: 1. Adjust the volumes of RNA samples to 4.5 µl and add: 2 µl 10× MOPS buffer 3.5 µl 37% formaldehyde 10 µl formamide Prepare the RNA molecular weight marker in the same manner.

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2. 3. 4. 5. 6. 7.

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Incubate at 60ºC for 15 min. Add 1/10 volume of 10× loading buffer. Apply RNA specimens to sample wells and electrophorese at 2.5 V/cm overnight. Remove the gel from the chamber and stain with ethidium bromide for 10 min. Rinse with water for at least 1 h to remove excess ethidium bromide and formaldehyde. Examine the gel on a ultraviolet transilluminator and photograph.

RNA Transfer RNA molecules fractionated on an agarose-formaldehyde gel should be transferred to a positively charged nylon membrane and immobilized on the membrane by baking or ultraviolet cross-linking before hybridization. Nylon membrane preparation: 1. Cut a piece of positively charged nylon membrane (Fisher Scientific) to the dimensions of the agarose gel and wet the membrane by carefully laying it on top of DEPC-treated water in a shallow tray. 2. Agitate the tray gently once the membrane is wet, then completely immerse the membrane in water. 3. Transfer the membrane to a second tray containing transfer buffer (20× SSC). Capillary blotting stack preparation: 1. Fill a tray with 1000 ml of transfer solution (20× SSC) and suspend a support, such as a glass plate, across the sides of the tray. 2. Wet two sheets of Whatman filter paper in 20× SSC. Then lay the sheets across the support with the ends dipping into 20× SSC. 3. Place the gel on top of the filter paper wicks carefully. Then place the membrane on top of the gel. Do not leave any portion of the gel exposed. To prevent the flow of buffer around the edge of the gel, place a strip of Parafilm along the edges of the gel. This acts as a barrier between the wicks and the absorbent material on top of the stack. 4. Wet three sheets of Whatman filter paper, cut to the size of the membrane, in 20× SSC and place them on top of the membrane. Roll out any trapped bubbles between the gel, membrane, and filter paper layers with a pipette. 5. Place a 20-cm-high stack of paper towels on top of the filter paper. Place a glass plate on top of the stack. Then place a 500-g weight (a large catalog) on top of the plate to evenly distribute the downward force. 6. Let RNA transfer from the gel to the membrane overnight. Then remove the paper towels and filter paper. 7. Lift the blot from the gel carefully with a pair of forceps. Rinse the blot with 5× SSC to remove loose agarose particles. RNA immobilization: 1. Fix the RNA to the membrane by baking in a vacuum oven (–25 in. vacuum mercury at 80ºC for 2 h) or by ultraviolet cross-linking (254 nm) in an ultraviolet transilluminator. 2. Stain the RNA on the membrane is stained at room temperature for 2 min in a solution that contains 0.5 M sodium acetate (pH 5.2) and 0.04% methylene blue. 3. Rinse the membrane in water for 5 to 10 min. Hybridization Analysis of RNA Blots The RNA blot is hybridized with a labeled DNA or RNA probe that is ideally 100 to 1000 bp long. Although hybridization using an RNA probe is more sensitive because of the greater stability of RNA–RNA hybrids, we routinely use radioactive DNA probes with satisfactory results. Hybridization is carried out in a solution containing formamide, which allows the hybridization to be

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performed at lower temperature reducing degradation of RNA on the membrane.10 Blocking reagents such as Denhardt’s solution and denatured salmon sperm DNA are also added to reduce nonspecific hybridization of the probe. After hybridization, the membrane is washed to remove nonspecifically bound probe molecules. The hybridized probes are detected by exposing the membrane to an X-ray film or a fluoroimage screen for various periods of time. Figure 13D.1 is an example of hybridization signals detected on RNA blots prepared from mouse eyes.12 Probes for tub and Tulp1 detected major transcripts at 6.3 and 2.4 kb, respectively. The membranes can be reused by stripping probes from the membrane without loss of RNA. This is particularly important when intensity of the bands in different lanes is to be compared on a blot. The same membrane should be stripped and hybridized with a probe for a stably expressed gene such as Gapd or Actb to demonstrate that the relative amount of RNA loaded in each lane of the gel is similar. Labeling of DNA probes by random priming: 1. Adjust DNA fragment to 50 ng in 34 µl H2O in a 1.5-ml Eppendorf tube. 2. Place DNA in boiling water for 5 min and cool the specimens on ice. 3. Centrifuge briefly. 4. Add: 5 ml 10× RP buffer 10× dNTP without dCTP Mix (1 mM each) 5 ml a-[32P]-CTP (3000 ci/mM) 5 ml Klenow Polymerase (NEB, 2 unit/µl) 1 ml 5. Mix well by vortexing. 6. Incubate at room temperature for 2 h. 7. Purify 32P-labeled DNA probe with a push column kit (Stratagene) according to the manufacturer’s instruction. 8. Determine the specific radioactive activity of 32P-labeled probe. Good labeling should give a specific activity >108 dpm/µg. Prehybridization and hybridization: 1. Prehybridize the membrane in hybridization solution (0.2 ml/cm2) at 45ºC for 2 to 4 h.

FIGURE 13D.1 Northern blot analysis of tub and Tulp1 mRNA in the eye. Poly(A)+ mRNA isolated from mouse eyes was subjected to electrophoresis, transferred onto nylon membranes and hybridized with 32P-labeled cDNA probes for tub (lane 1) and Tulp1 (lane 2). RNA size markers are shown on the left. Lane 1 was exposed for 3 h and lane 2 was exposed for 7 min.

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2. Hybridize the membrane with 32P-labeled nucleotide probe (106 cpm/ml) in hybridization solution at 45ºC overnight. Washing: 1. First wash with 2× SSC/0.1% SDS at room temperature for 30 min. 2. Then wash twice with 2× SSC/0.1% SDS at 65ºC for 30 min. 3. Further wash with 0.2× SSC/0.1% SDS at 65ºC for 30 min. X-ray film exposure: 1. Expose the membrane to Kodak T-Mat G/RA X-ray film or fluoroimage screen for various periods of time. RNA blot stripping and rehybridization: 1. Prepare stripping solution. 2. Gently place the RNA blot in a glass baking dish. Take care not to dry the membrane between hybridization and stripping. 3. Pour boiling stripping solution onto the membrane. Allow the solution to cool to room temperature. 4. Seal the blot in a plastic bag. Expose the blot to film for normal exposure time to assure the removal of 32P-probe. If the probe has not been removed from the blot, repeat the stripping procedures. When the probe has been removed, the blot can be hybridized with a different 32P-labeled probe. Reagents and Solutions 10× MOPS buffer 0.2 M MOPS (3-(N-morpholino)-propanesufonic acid), pH 7.0 0.1 M sodium acetate 0.01 M EDTA 10× loading buffer 50% glycerol 1 mM EDTA 0.25% bromophenol blue 0.25% xylene cyanol FF 10× RP buffer 0.5 M Tris-HCl (pH 7.5) 0.1 M MgCl2 10 mM DTT 0.5 mg/ml BSA 0.05 A260/µl pd (N)6 (Pharmacia LKB 27-2166-01) Hybridization solution 1 M HEPES (pH 7.0) 100× Denhardt’s solution 20× SSC Salmon sperm DNA (10 mg/ml) 10% SDS Formamide (super pure grade) Add H2O to

50 ml 10 ml 300 ml 15 ml 10 ml 500 ml 1000 ml

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100× Denhardt’s solution Ficoll 400 10 g Polyvinylpyrrolidone 10 g Bovine serum albumin 10 g Add H2O to 500 ml Filter sterilize the solution and store at –20ºC. 20× SSC NaCl Na3citrate × H2O Add H2O to Adjust pH to 7.0 with 1 M HCl Add H2O to

175.3 g (3 M) 88.2 g (0.3 M) 800 ml 1l

Salmon sperm DNA (Sigma type III sodium salt) Dissolve DNA in water (10 mg/ml) at room temperature for 2 to 4 h. Adjust NaCl concentration to 0.1 M final concentration. Extract once with Tris-HCl buffer (pH 8.0)–saturated phenol and once with phenol:chloroform (1:1). Recover the aqueous phase (which contains the DNA) and shear the DNA by passing it 12 times rapidly through an 18-gauge hypodermic needle. Precipitate the DNA with equal volume of isopropanol. Recover the DNA precipitate by centrifugation and redissolve the DNA in water (10 mg/ml). Denature DNA by boiling for 15 min and store at –20ºC. Stripping Solution 0.5% (w/v) SDS

IN SITU HYBRIDIZATION In situ hybridization is a method of detecting mRNA within tissues and cells using labeled complementary nucleotide sequences. It reveals the spatial pattern of expression of a gene in a heterogeneous cell population, permitting determination of whether a gene is expressed uniformly in all cell types within a tissue or only in certain cell types. As an example, Figure 13D.2 shows mouse retina hybridized with 33P-labeled probes for tub and Tulp1 RNAs.12 The tub mRNA is expressed throughout the retina, whereas Tulp1 mRNA is expressed only in photoreceptor cells. Fixation and Permeability of Tissue The tissue used for in situ hybridization should be fixed or frozen as soon as possible after harvesting to avoid RNA degradation. Fixation of the tissue is recommended prior to freezing to maintain tissue morphology. Paraformaldehyde (PFA) is the most widely used fixative for in situ hybridization. After fixation, the tissue is frozen and processed for cryosectioning or processed for paraffin embedding/sectioning. Sections should be treated with a proteinase before hybridization to increase permeability of the tissue and allow for penetration of the probe in the tissue sections. This is because the sequence to be detected may be present at a low concentration or may be masked or protected within cells and cellular structures, such as ribosomes. Probe Labeling: Radioactive or Nonradioactive Both radioactive and nonradioactive probes have been used successfully for in situ hybridization. The method for probe labeling should be chosen based on the tissue/probes to be treated and on the

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FIGURE 13D.2 Detection of tub and Tulp1 mRNA in the retina by in situ hybridization. A and B. Brightfield Sections C and D. Corresponding dark-field sections. tub expression was detected throughout the retina (C) and Tulp1 was detected exclusively in photoreceptor cells (D). GCL = ganglion cell layer, INL = inner nuclear layer, ONL = outer nuclear layer.

advantages and disadvantages of each method. Hybridization using radioactive probes provides a more sensitive indication of hybridization,13 and quantification of signal is possible by counting grains.14 However, radioactive labeling requires greater laboratory safety measures and may require relatively long exposure times (weeks to months) to obtain useful results. The shelf life of a radioactive probe is defined by the half-life of the isotope used for labeling. With regard to the choice of isotopes for labeling, 33P has become more widely used because of its relative safety, reasonably shorter exposure time, and good resolution.15 Nonradioactive probes have the advantages of safety and speed with results within 3 to 4 days. It is suitable when exact, cell-specific localization of signal is desired, since the signal is viewed under bright-field conditions. In addition, nonradioactive probes can be stored for an extended time (months to years). The disadvantages of nonradioactive probes are that low-level labeling is difficult to distinguish from background and quantification is difficult.16 The most widely used nonradioactive label, digoxigenin (DIG),17 is of plant origin, and produces low, nonspecific staining on animal tissue sections. The hybridization signals with DIG-labeled riboprobes are detected by immune reaction with alkaline phosphatase-conjugated anti-DIG antibodies followed by enzyme substrate reaction.18 DIG-labeled probes provide the same degree of sensitivity as radioactive probes for signal detection19 and are efficient alternatives to radiolabeled probes. The success of in situ hybridization depends largely on the quality of the probe. Specificity of the probe should be evaluated by Northern hybridization using a blot made from the tissue of interest before it is used for in situ hybridization. The result will provide an indication of how specific the probe is and how abundant the target mRNA is in the tissue.

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Single-Stranded RNA Probes The use of single-stranded RNA probes is preferred to cDNA probes for in situ hybridization because of the high affinity of RNA–RNA hybrids.20 It is also an advantage that nonspecifically bound probes can be removed by RNase A treatment during the posthybridization washing to reduce the background. To generate an RNA probe, the cDNA fragment is subcloned into a vector that has RNA polymerase promotors (T7, T3, or SP6) on both sides of the multiple cloning site (Figure 13D.3). This serves as a template for the RNA polymerase reaction. The template is linearized with a restriction enzyme at one end of the insert to limit the length of the probe produced. An antisense probe is generated by in vitro transcription using an appropriate RNA polymerase (T3 polymerase in Figure 13D.3). Antisense probes are complementary to target mRNAs in the tissue and should hybridize to them. Using the same template, a sense probe should also be generated. Sense probes are used as internal negative controls for hybridization since they are identical to mRNAs in the tissue and do not hybridize to them. Conditions of Hybridization and Washing The degree of specificity of hybridization depends on numerous factors such as the sequence of the probe, temperature, concentration of a denaturant such as formamide, and concentration of salt in the hybridization buffer.21 Although only probes with high homology to the target sequence hybridize under conditions of high stringency, probes with numerous mismatches can form hybrids under conditions of low stringency. In most protocols, hybridization buffers contain 50% formamide and 2× SSC (standard saline citrate-salt solution). Dextran sulfate is also added to increase hybridization efficiency. Hybridization is carried out from 37 to 60°C depending on the stringency desired. Washing steps are performed to reduce nonspecific binding of the probes. The sections are incubated in decreasing concentrations of SSC with increasing stringency. Conditions for hybridization and washing can be modified depending on the results obtained.

cDNA HindIII

5’

3’

T7

BamHI T3

pBluescript vector HindIII cut

BamHI cut

antisense RNA probe

sense RNA probe T3

T7

T3

T7

FIGURE 13D.3 Diagrammatic representation of RNA probe generation. A cDNA fragment is ligated into a plasmid, which is linearized at either end of the insert. Single-stranded antisense and sense RNA probes are generated by in vitro transcription using appropriate RNA polymerases (T3 for antisense and T7 for sense probes for this construct).

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Visualization of the Signal After hybridization and washing, the signal is visualized depending on the probe system used. If radioactive probes are used, the signal is detected by making autoradiograms with liquid emulsions. The signal visualized under dark-field conditions is compared with the bright-field image to demonstrate location of the signal within the tissue. If DIG-labeled nonradioactive probes are used, the signal is detected by an immune reaction using an anti-DIG antibody conjugated with alkaline phosphatase. The substrates for alkaline phosphatase, NBT (4-nitro blue tetrazolium chloride) and BCIP (5-bromo-4-choro-3-indolyl-phosphate), are added to detect antibody binding. Protocols for in Situ Hybridization The procedures employed successfully in our laboratory using paraffin-embedded tissues and DIGlabeled RNA probes will serve as an example of protocols for in situ hybridization. The protocol for in situ hybridization using radioactive probes is basically similar to this protocol until the washing step after hybridization except that the probe is labeled with 33P instead of DIG.12,13,15 After the washing step, the signal is detected by autoradiography using liquid emulsion. Preparation of paraffin-embedded tissue blocks: 1. Fix dissected tissues immediately in 4% PFA in 0.1 M phosphate buffer, pH 7.4 for 24 h at 4°C. For larger tissues, e.g., whole embryos at E16.5, extend fixation time to 48 h with two changes of fixative solution. 2. The next day, wash the tissues twice with PBST (PBST = PBS + 0.1% Tween-20). 3. Dehydrate the tissues through a graded series of alcohol prepared with DEPC-treated water and embed in paraffin. The paraffin blocks can be stored at 4°C for extended periods of time. We have been able to obtain excellent hybridization signals with paraffin blocks stored for more than 1 year. Sectioning: 1. Use a microtome set to 5-micrometer sections to produce “ribbons” of consecutive sections. 2. Transfer the sections to the preheated DEPC-treated water bath to unroll the paraffin ribbons. 3. Mount the sections onto Fisher brand ProbeOn Plus microscope slides (Fisher Scientific, Pittsburgh, PA), which are charged and precleaned. 4. Incubate slides overnight at 65ºC to melt the paraffin and to allow the tissue to settle directly onto the slide surface. Preparation of the DIG-labeled RNA probe: The labeling mixes as well as all antibodies can be purchased from Roche. All conditions and solutions should be absolutely RNase-free. 1. Linearize the plasmid and verify the completion of enzyme digestion by agarose gel electrophoresis. 2. Extract the linearized plasmid DNA once with phenol, once with phenol:choloform: isoamylalcohol (25:24:1), and once with chloroform:isoamylalcohol (24:1), and precipitate with ethanol. 3. Resuspend the linearized plasmid DNA in DEPC-water. 4. Determine the DNA concentration with spectrophotometer at 260 nm. 5. Set up transcription reaction: 1 mg linearized plasmid DNA 6 µl 10× transcription buffer (Boehringer Mannheim, Indianapolis, IN) 6 µl labeling mix (Boehringer Mannheim) 1.5 µl RNAsin (40 unit/microliter, Promega, Madison, WI)

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6. 7. 8.

9. 10. 11. 12.

6 µl RNA polymerase (20 unit/ml) (SP6, T3 or T7, Boehringer Mannheim). Add DEPC-treated water to 60 µl and incubate at 37°C for 2 h. Add 6 µl RQI RNAse-free DNAse I (10 unit/ml) (Promega) and incubate at 37ºC for 15 min. Examine the size of the synthesized riboprobe by electrophoresis in 1% agarose gel (2 µl of the probe ). The RNA should appear as a single band with little degraded product. Dilute the reaction mixture with an equal volume of DEPC-treated water. The riboprobe can be partially hydrolyzed by incubating the specimens in 0.3 M NaHCO3 pH 10.2 (add equal volume of 0.6 M NaHCO3, at pH.10.2, to the sample and incubate at 60ºC for 20 to 40 min). The size of the riboprobe should be analyzed at 10-min intervals by agarose gel electrophoresis. The ideal size of RNA probes is about 250 to 500 bases. Precipitate the RNA probe by adding 40 µl of 3 M sodium acetate (pH 5.2) and 1 ml of 100% ethanol to the mixture. Centrifuge, then resuspend the RNA probe in DEPC-treated water. Determine the concentration of the RNA probe by spectrophotometer. Aliquot the RNA probe (10 µl/tube) and store at –70ºC. (The probe can be stored for 1 to 2 years if repeated freeze/thaw is avoided.)

Pretreatment of sections: 1. Deparaffinize and rehydrate sections through graded series of alcohol to water (DEPCtreated). 2. Wash sections in PBS for 5 min. 3. Fix in 4% PFA in PBS at room temperature for 5 min. 4. Wash in PBS for 5 min (three times). 5. Proteinase K digestion: Treat with Proteinase K (Sigma, 50 mg/ml) at 37ºC for 10 min and wash three times with PBS, 2 min each. 6. Fix in 4% PFA in PBS at room temperature for 10 min and wash three times with PBS, 2 min each. 7. Acetic anhydride blocking. Treat the sections with glycine solution (10 mg/ml) in PBS at room temperature for 5 min, then dip in 0.1 M triethanolamine/PBS (pH 8.0) for 10 min, and further incubate in a freshly made 0.25% (v/v) acetic anhydride in 0.1 M triethanolamine/PBS (pH 8.0) for 5 min. This treatment decreases nonspecific binding of the probe. 8. Dehydrate the sections through graded series of alcohol and air dry for 2 h at room temperature. Prehybridization: Cover the tissue sections with 1× prehybridization buffer/50% formamide solution (30 to 50 µl/tissue section) in a moist chamber equilibrated with 50% formamide at 45ºC for 2 to 4 h. Hybridization: Remove the prehybridization solution using a piece of Kimwipe EX-L (Kimberly-Clark) and cover the tissue sections with 1× hybridization buffer/50% formamide solution containing 2 to 10 µg/ml RNA probe. Hybridization is done at 45ºC in a moist chamber overnight. Washing: 1. Prewarm 0.5× SSC and 1 mM EDTA solution to 55 to 60ºC and drop it on the slides to remove RNA probes. Never allow the slides to dry until after all washing steps are complete. 2. Wash in 0.5× SSC + 1 mM EDTA at room temperature for 10 min (three times).

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3. Wash in 0.5× SSC + 50% formamide + 1 mM EDTA at room temperature for 30 min. 4. Incubate in solution containing RNase A (20 µg/ml) at 37ºC for 30 min to remove nonspecifically bound RNA probes. 5. Wash further in 0.5× SSC + 1 mM EDTA at room temperature for 10 min (3 times). Anti-Digoxigenin antibody reaction: 1. Incubate sections with a blocking solution (30 to 50 µl/tissue section) containing 3% skim milk, 5% normal goat serum in 1× Tris-saline at room temperature in a moist chamber for 60 min. 2. Remove the blocking solution with a Kimwipe EX-L and incubate sections with diluted (1:2000) anti-Dig Fab alkaline phosphatase coupled antibody (Roche) in 1× Tris-saline + 3% skim milk overnight at 4ºC in a moist chamber equilibrated with water. Washing: 1. Wash in Tris-saline for 20 min (three times). 2. Incubate with AP buffer (pH 9.5) for 5 min. Color development: 1. Prepare the color reaction solution. To 1 ml AP buffer add: 4.5 µl NBT (75 mg/ml) (Boehringer Mannheim), in 70% dimethylformamide 3.5 µl BCIP (50 mg/ml) (Boehringer Mannheim), in 100% dimethylformamide 0.24 mg/ml levamisole 2. Apply 100 to 200 µl reaction solution per slide and incubate at room temperature for 2 to 8 h in the dark. Mounting: 1. Rinse sections with water. 2. Air-dry sections overnight. 3. Counterstain sections with 0.5% neutral red for 8 min. 4. Dehydrate in a graded series of ethanol. (Note: Some signal is lost during this process, so dehydrate only 5 min at each step.) 5. Mount the sections with Permount (Fisher Scientific). Reagents and Solutions 2× Prehybridization buffer stock 8.0 ml 20× STE 0.4 ml 100× Denhardt’s solution 2.0 ml 10 mg/ml sheared, denatured salmon sperm DNA 2.4 ml 10 mg/ml yeast tRNA 7.2 ml H2O/DEPC 20× STE 3 M NaCl 0.6 M Tris-HCl (pH 8.0) 0.04 M EDTA 2× Hybridization buffer Dextran sulfate 20× STE 100× Denhardt’s solution 10 mg/ml sheared, denatured salmon sperm DNA 10 mg/ml yeast tRNA Add water to 20 ml total.

4g 8.0 ml 0.4 ml 0.4 ml 0.4 ml

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Tris-buffered saline 1 M Tris-HCl (pH 7.4) 1.5 M NaCl H2O

100 ml 100 ml 800 ml

AP buffer 100 mM Tris pH 9.5 100 mM NaCl 50 mM MgCl2 (Note: Tris pH 9.5 must be stored as frozen aliquots as the solution will become pH < 9.0 if stored at room temperature.)

REFERENCES 1. Ying, S. et al., Characterization and expression of the mouse lumican gene, J. Biol. Chem., 272: 30306, 1997. 2. Saika, S. et al., Role of lumican in the corneal epithelium during wound healing, J. Biol. Chem., 275: 2607, 2000. 3. Liu, C.Y. et al., The cloning of mouse keratocan cDNA and genomic DNA and the characterization of its expression during eye development, J. Biol. Chem., 273: 22584, 1998. 4. Schena, M. et al., Quantitative monitoring of gene expression patterns with a complementary DNA microarray, Science, 270: 467, 1995. 5. Gibson, U.E., Heid, C.A., and Williams, P.M., A novel method for real time quantitative RT-PCR, Genome Res., 6:995, 1996. 6. Heid, C.A. et al., Real time quantitative PCR, Genome Res., 6:986, 1996. 7. Turnbow, M.A. and Garner, C.W., Ribonuclease protection assay: use of biotinylated probes for the detection of two messenger RNAs, Biotechniques, 15:267, 1993. 8. Chomczynski, P. and Sacchi, N., Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction, Anal. Biochem., 162:156, 1987. 9. Chomczynski, P. and Mackey, K., Short technical reports. Modification of the TRI reagent procedure for isolation of RNA from polysaccharide- and proteoglycan-rich sources, Biotechniques, 19:942, 1995. 10. Brown, T. and Mackey, K., Analysis of RNA by Northern and slot blot hybridization, in Current Protocols in Molecular Biology, Ausubel, F. M. et al., Eds., John Wiley & Sons, New York, 1997, chap. 4.9. 11. Lehrach, H. et al., RNA molecular weight determinations by gel electrophoresis under denaturing conditions, a critical reexamination, Biochemistry, 16:4743, 1977. 12. Ikeda, S. et al., Cell-specific expression of tubby gene family members (tub, Tulp1, 2, and 3) in the retina, Invest. Ophthalmol. Vis. Sci., 40:2706, 1999. 13. Hoover, S.B. et al., In situ hybridization techniques for studying gene expression in the genetically altered mouse, in Pathology of Genetically Engineered Mice, Ward, J. et al., Eds., University of Iowa Press, Ames, 2000, chap. 2. 14. Aldridge, J., Quantification of grains in in situ hybridization, in In Situ Hybridization, Polak, J.M. and McGee, J.O., Eds., Oxford University Press, New York, 1998, chap. 2. 15. Bisucci, T., Hewitson, T.D., and Darby, I.A., cRNA probes: comparison of isotopic and nonisotopic detection methods, Meth. Mol. Biol., 123:291, 2000. 16. Jilbert, A.R., In situ hybridization protocols for detection of viral DNA using radioactive and nonradioactive DNA probes, Meth. Mol. Biol., 123:177, 2000. 17. Komminoth, P., Digoxigenin as an alternative probe labeling for in situ hybridization, Diagn. Mol. Pathol., 1:142, 1992. 18. Fleming, K.A. et al., Optimization of non-isotopic in situ hybridization on formalin-fixed, paraffin-embedded material using digoxigenin-labelled probes and transgenic tissues, J. Pathol., 167:9, 1992. 19. Komminoth, P. et al., Comparison of 35S- and digoxigenin-labeled RNA and oligonucleotide probes for in situ hybridization. Expression of mRNA of the seminal vesicle secretion protein II and androgen receptor genes in the rat prostate, Histochemistry, 98:217, 1992.

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20. Sunday, M.E., The design and generation of probes for in situ hybridization, in In Situ Hybridization, Polak, J.M. and McGee, J.O., Eds., Oxford University Press, New York, 1998, chap. 2. 21. Hofler, H., Mueller, J., and Werner, M., Principles of in situ hybridization, in In Situ Hybridization, Polak, J.M. and McGee, J.O., Eds., Oxford University Press, New York, 1998, chap. 1.

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Other Techniques Gregory Martin, Richard S. Smith, Simon W. M. John, Olga V. Savinova, Steven Nusinowitz, William H. Ridder III, and John R. Heckenlively

CONTENTS A. Confocal Microscopy, Morphometrics, and Cell Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . .300 Gregory Martin and Richard S. Smith Confocal Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .300 Principles of Confocal Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .300 Multi-Dimensional Confocal Data Sets . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .302 Utility of Confocal Microscopy in Thick and Living Tissues . . . . . . . . . . . . . . . . . .306 Morphometrics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .307 Simple Morphometrics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .307 Quantitative Morphometrics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .308 Cell Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .309 Ki-67 Antigen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .309 Proliferating Cell Nuclear Antigen (PCNA) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .310 Bromodeoxyuridine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .310 Radioactive Labeling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .311 Mitotic Rates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .311 General Problems of Interpretation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .311 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .312 B. Intraocular Pressure Measurement in Mice: Technical Aspects . . . . . . . . . . . . . . . . . . . . . . . .313 Simon W. M. John and Olga V. Savinova Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .313 Important Dimensions and Volumes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .314 Measurement Procedure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .314 Anesthesia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .318 Stress and Other Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .319 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .319 C. Electrophysiological Testing of the Mouse Visual System . . . . . . . . . . . . . . . . . . . . . . . . . . . .320 Steven Nusinowitz, William H. Ridder III, and John R. Heckenlively Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .320 The Full-Field Electroretinogram . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .320 Basic ERG Recording Technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .321 Isolating Rod- and Cone-Mediated Responses . . . . . . . . . . . . . . . . . . . . . . . . . . . . .322 Accuracy of the ERG . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .324 Examples of ERG Patterns in Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .325 Factors Affecting the ERG . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .326 Anesthetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .327 Mouse Body Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .328 Pupil Size . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .328 0-8493-0864-X/02/$0.00+$1.50 © 2002 by CRC Press LLC

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Dark and Light Adaptation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .328 Specialized ERG Recording Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .329 a-Wave Analyses: Studies of Activation and Inactivation Steps of Phototransduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .330 The Multifocal Electroretinogram (MERG) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .332 Visual Evoked Cortical Potentials (VECP) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .333 The Mouse Visual Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .334 VECP Stimuli . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .334 Flash VECP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .335 Pattern VECP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .336 Multifocal VECP (MVECP) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .338 General Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .340 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .340

A. CONFOCAL MICROSCOPY, MORPHOMETRICS, AND CELL KINETICS Gregory Martin and Richard S. Smith

CONFOCAL MICROSCOPY Confocal microscopy is a technique particularly well suited to investigations of the mouse eye, an organ in which three-dimensional structure and function are intimately related. Selecting the appropriate imaging technology based on the scientific questions that are being addressed is key to an analysis of structure–function relationships. With careful selection of instrumentation and experimental design, confocal images of structures of interest can be combined with information about the function of living tissues. Most life scientists have at one time or another looked at a relatively thick specimen with a microscope and noticed that only one plane within the specimen was in sharp focus. Regions of the specimen above and below this focal plane appear as out-of-focus blur. Worse still, these out-of-focus regions degrade the image formed at the focal plane, reducing both apparent specimen contrast and resolution. From the aspect of image formation, signal emitted from above and below the focal plane is collected and delivered to the detector—an observer or camera—where it contributes to and degrades the desired image from the focal plane. The objective of confocal microscopy is to exclude the out-of-focus information from the detector and collect an image only from the focal plane. Principles of Confocal Microscopy Confocal microscopy is a robust and extremely useful technique that was initially developed and patented by Marvin Minsky in the late 1950s.1 A thorough discussion of the theory and practice of confocal microscopy is beyond the scope of this chapter. Interested readers can obtain further information from several excellent texts on the subject2-4 including a treatise dealing specifically with ocular morphology.5 Although confocal microscope instrumentation can be very complex, the basic principle is not and any discussion must start with fluorescence microscopy. The majority of applications for confocal microscopy in biology utilize fluorescent techniques. For an introduction to fluorescence microscopy and the wide variety of probes available, the reader is referred to Mason.6 Briefly, fluorescence microscopy utilizes either labeled probes with an affinity for a specific cellular component or dyes that change their fluorescent characteristics in response to cellular events. Use of multiple probes labeled with different fluorescent compounds (fluorophores) provides a means to examine the fine structure and interrelationships of multiple tissue components or events. Excitation and emission wavelengths are specific to the various fluorophores. High-intensity

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excitation energy is provided by an arc lamp and filters or, as is most often the case for confocal microscopes, by lasers. After collection by the objective lens, the emitted fluorescent wavelengths are then specifically selected using optical filter sets to form an image. To eliminate contribution of out-of-focus planes to the final image in a confocal microscope, a variable pinhole is placed in front of the detector. At this plane light rays from the focal plane of the objective (that is, from the in-focus part of the specimen) are converging while light rays originating from above or below the focal plane are diverging. Thus, by setting the pinhole to an appropriate size for the objective being used, light from the focal plane will pass through the pinhole and be detected— used to form an image— while light from above or below the focal plane will be rejected. The rejection of this out-of-focus “blur” has several advantages. The most immediately obvious is an increase in image contrast and apparent resolution in the plane of the image, referred to as the xy plane. The practical effect is to produce an “optical section” from within the three-dimensional specimen (Figure 14A.1).* Optical sectioning provides one of the real benefits of confocal microscopy: the ability to better image complex structures that vary along the “depth” dimension as one looks down along the optical axis of the microscope. Images are data when investigating the interrelationship of structure and function. It is particularly true with the confocal microscope, which can be considered as much a sampling instrument as an imaging system. In the laser scanning confocal microscope mirrors are used to scan a point of laser excitation light over the specimen. The detector behind the pinhole is a photomultiplier tube that produces a signal in response to the number of photons striking it. Thus, the image is formed in a point-by-point manner with each point in the image representing the intensity of the fluorescent signal collected at that point in the specimen. By using lasers of different wavelengths for excitation and carefully chosen filters to separate different emitted wavelengths, multiple labels can be imaged simultaneously. This means that the confocal microscope is particularly well suited for multiple-labeling studies. Because the different fluorescent signals are collected at the same time,

FIGURE 14A.1 Wide field vs. confocal imaging. Wide field (A) and confocal (B) imaging of the same region of an isolated mouse retina. The retina was labeled with FITC-phalloidin, which specifically binds to f-actin. The brightly labeled structure running diagonally through the images is a blood vessel. The images were obtained with the same objective lens, and the image acquisition parameters of each instrument were set to yield images with a similar range of brightness values. * All images in this chapter are best viewed using the CD. Some images are available only on the CD.

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FIGURE 14A.2 Multilabel confocal imaging. Paraffin section of mouse retina labeled for DNA with YOYO-1 (A) and for fragmented DNA with BODIPY-based in situ end-labeling of fragmented DNA (B). Both fluorescent images and the DIC image were collected simultaneously. An apoptotic nucleus containing both highly condensed and fragmented DNA is indicated by the arrowhead. The color version of this plate also contains an overlay of the two fluorescent images (D), and the fluorescent overlay combined with the DIC image (E). Viewing of D and E is possible only on the CD.

through the same optical path, there is no registration shift between the images representing the individual labels. An example of multichannel imaging is shown in Figure 14A.2. In this case, the goal was to identify apoptotic cells in paraffin sections of the retina. The DNA-binding dye YOYO1 was used to label chromatin, permitting visualization of nuclei with highly condensed chromatin. In addition, a fluorescence-based in situ end-labeling of fragmented DNA7 was performed to highlight nuclei containing fragmented DNA (arrowhead, Figure 14A.2B). A differential interference contrast (DIC) image (Figure 14A.2C) was also collected to provide morphological data. Each of the three images provides information from a different technique, each relevant to the identification of apoptotic cells. Multi-Dimensional Confocal Data Sets Collecting optical sections at intervals through the depth of the specimen provides a means of sampling the three-dimensional (3D) distribution of fluorescent probes. After collection of a “stack” of images with the confocal microscope, software is required to visualize the resulting 3D data set. Most instruments include some capability for manipulating and visualizing stacks of images as part of the software that runs the instrument. More thorough treatments of 3D image processing and analysis than are within the scope of this chapter are readily available.5.8,9 For the purposes of this discussion, the microvasculature within the retina of a mouse that was perfused with fluorescent dextran10 provides an excellent example of 3D imaging with the confocal microscope. A single image stack will be used in the following examples to illustrate some basic methods of examining the three-dimensional structure of tissue. The original image stack was collected with a 10× lens, and consists of 54 images collected at 2-µm intervals through the entire thickness of a whole-mounted retina. Perhaps the most straightforward method for analyzing image stacks is by displaying the individual images as a gallery. Selected images from the original data set are arranged as a gallery in Figure 14A.3. In this case, depicting all 54 planes from the stack is unnecessary because adjacent images are too similar, so every fourth image is shown, depicting optical sections from 8 µm intervals. The effect of the optical sectioning can clearly be seen. Computer software can be used to display the individual images in succession, allowing one to obtain a sense for how the distribution of the label changes with depth. The individual images can be made to play as a movie, giving the effect of focusing through the specimen.

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FIGURE 14A.3 Series of confocal images from an isolated mouse retina. Optical sections of fluorescently labeled vasculature were collected at 2-µm intervals through the entire thickness of the preparation. Individual images from the data set were selected at 8-µm intervals for display.

Another common method for visualizing 3D data sets is by making a projection image. A projection image is just that; image data from all of the individual images of a stack are projected down into a single image. Most confocal systems contain software with the ability to generate projection images. Using the same retinal vascularization image stack as an example, a projection image is shown in Figure 14A.4. Note that vascular elements present in either the top (Figure 14A.3A) or the bottom (Figure 14A.3L) of the stack are both evident in the projected image (Figure 14A.4). The projection image can be very useful for visualizing the distribution of label in the xy plane without regard to depth (the z dimension). Useful spatial information can also be obtained by judicious combinations of projected images of a structural marker to show overall morphology and a single plane of some other label of interest. Projection images form the basis for other more advanced 3D visualization techniques. Appropriate computer software can, in effect, tilt the original stack of images to the left or right prior to creating a projection. The resulting projection images can be treated as single “views” through the stack at the specified angle. By creating two projection images of the stack at angles slightly to the left and right of center and using optical devices (such as special glasses) to deliver those images to the left and right eye (respectively) of the viewer provides an impression of three dimensionality. Figure 14A.5 is such a stereo projection from the same data set used for Figures 14A.3 and 14A.4. Stereo pairs displayed side by side can be viewed with stereo viewers, although some people can merge the two images without the aid of a viewer. Two color stereo pairs such as shown on the CD can be viewed easily with inexpensive viewers containing colored lenses. The color technique is more straightforward for projection, but lends itself less well to mutilabel (i.e., multicolor) image sets. These stereo pairs are easily created on the computer and very useful for evaluating the 3D distribution of label. As

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with the use of multilabel image data for projection images, a structural marker could be displayed as a stereo view, with another marker of interest overlaid as individual image planes. The next level of displaying 3D confocal image data is to create a series of projections, each at a different angle of rotation of the original stack of images. By displaying this series of projections in sequence, an animation can be produced of the 3D image set turning in space. These sorts of computer-generated animations are useful tools for display and evaluation of confocal data sets. Much more advanced techniques are available, depending on the computer programs used, to analyze, or

FIGURE 14A.4 A projection image constructed from the series of confocal images depicted in Figure 14A.3.

FIGURE 14A.5 A stereo projection constructed from the series of confocal images depicted in Figure 14A.3. The pair of black-and-white images should be viewed with stereo viewers, while the color version should be viewed with red/green glasses (available only on the CD).

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render, the data set into sophisticated 3D reconstructions as well as to perform 3D quantification measuring distance, volume, or surface area. Often a complete 3D data set is not needed. Because the image is created point by point by scanning the excitation beam within the specimen, one can collect an image from a plane other than the standard xy plane. By moving the specimen in the z-dimension, just as for collecting an image stack, but scanning the excitation beam along a single line in the x dimension, an xz image is generated. An example of xz imaging is shown in Figure 14A.6. In this case the specimen is an isolated retina immunolabeled for blue opsin, a molecule found in the outer segments of blue-sensitive cone cells within the photoreceptor layer of the retina. The immunofluorescence image is shown in Figure 14A.6A, pseudocolored green in the color version of the figure. It is an xy image at the level of the cone outer segments. Figure 14A.6B, shown in red in the color version, is the image of a DNA counterstain collected 24 µm deeper within the retina, within the outer nuclear layer. Two xz images are shown in Figures 14A.6C and D, collected at the planes indicated in Figure 14A.6A. Although both

FIGURE 14A.6 Confocal XZ imaging. An isolated mouse retina was labeled with an antibody against blue opsin and counterstained with the DNA-specific dye TO-PRO 3. An xy image of the blue opsin immunolabeling is shown at the level of the outer segments A. An xy image of the nuclear counterstain, collected 24 µm deeper within the outer nuclear layer, is shown in B. XZ images, collected at the planes indicated in (A) are shown (C and D) as color overlays in the color version of the figure, which is available on the CD.

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labels are shown in gray scale for the black-and-white version of the figure, the location and pattern of the two labels (blue opsin and DNA) are different enough to provide a useful combined image. Utility of Confocal Microscopy in Thick and Living Tissues There are, of course, limits to how deep useful images can be gathered with the confocal microscope. The retinas used as examples above were 100 to 150 µm thick and the vascular labeling technique produces a bright label with very little background. Imaging such a specimen at low magnification provides useful images through the entire thickness of the retina. However, this is the exception rather than the rule. More often the situation is less optimal: the label is weaker, the background higher, and/or the appropriate magnification is higher. These and many other tissue and instrument factors can severely limit the depth from which useful images can be obtained.11 Figure 14A.7 demonstrates a more typical example of the changes in image quality with depth. The specimen is the same isolated retina immunolabeled for blue opsin and DNA-counterstained as was used for the images in Figure 14A.6. In this case, the margin of the retina has rolled up, so that the confocal is imaging a cut edge of the retina. Figure 14A.7A was collected at the surface of the cut edge of the retina, as evidenced by the crack observed across the specimen. Note that the subcellular distribution of blue opsin is clearly evident. Imaging just 10 µm deeper within the tissue (Figure 14A.7B) gives a slight loss in apparent resolution. The effect is greater 20 µm into the specimen (Figure 14A.7C). At 30 µm depth (Figure 14A.7D), the image has degraded further but may still be useful. Images collected deeper (Figures 14A.7E and F) have a significantly lower signal to noise ratio, and may be unusable. One of the most exciting applications of confocal microscopy is its use to study the 3D organization of living tissues. In the field of eye research the clinicians have taken the lead in this area. Several reviews12-15 are available that detail the use of confocal instrumentation for studying the threedimensional organization and abnormalities in the living eye. Most of these techniques should be

FIGURE 14A.7 Loss of resolution with depth. An isolated mouse retina was labeled with an antibody against blue opsin and counterstained with the DNA-specific dye TO-PRO 3. Imaging the surface of the cut edge of the retina gives the best image (A). Subsequent images (B to F), taken at 10-µm intervals, show a progressive increase in signal-to-noise ratio and a loss in apparent resolution.

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applicable to the mouse eye. Examples of the use of confocal microscopy for in vivo analysis in research-oriented settings include studies of neovascularization,16 corneal fibrosis,17 and photoreceptor development using green fluorescent protein.18 An entirely different class of fluorescent probes can be used in living tissues. Such probes are generally not meant to bind to a particular cellular element, but alter their fluorescent characteristics in response to their environment. These probes allow visualization and measurement of physiological events at the cellular level. The most familiar of these probes are those that act as indicators of intracellular ion concentrations.19 Indicators of intracellular Ca2+ concentration have been used to examine signaling mechanisms ex vivo (living, but isolated, tissue) imaging of ocular ciliary epithelium20 and in vitro imaging of primary cultures of corneal epithelium.21 Data sets from living specimens will often contain the additional dimension of time (t) as either xy-t series (imaging a single plane over time) or, more rarely, xyz-t series (imaging a multi-plane volume over time). It should be kept in mind the time “resolution” that is relevant to the study. How often is an event occurring and how often is the specimen being sampled? If the experimental needs so require, and the specimen conditions are appropriate, more rapid sampling can often be done by sampling in fewer dimensions (a line or a point) over the time during the course of an experiment.

MORPHOMETRICS* The term morphometrics refers to a variety of techniques designed to gather data from images for the measurement of biological structures of interest. The goal is to quantitate measurable differences in cells and tissues between populations of animals or experimental treatments. These differences often form the basis for understanding structure–function relationships. In some instances, simple techniques are capable of providing useful results that clearly demonstrate an effect. As more sophisticated computer software and hardware become available, increasingly complex measurements, such as 3D analysis and estimation of total numbers of specific structures in a tissue have become possible. Simple Morphometrics In many experimental designs there is a need to measure the differences in tissue layer thickness between control and mutant mice to determine if the visually observed difference is statistically significant. To make such measurements meaningful, tissue boundaries must be well defined. In the eye, however, thickness measurements are complicated by normal variations in the size of the structure. As an example, the peripheral retina is less than half the thickness of the retina adjacent to the optic nerve. When measurements are made, both the location and the extent of the region measured become part of the protocol. An example of a set of guidelines for measuring retina might include: 1. A field beginning at the edge of the optic nerve and extending temporally to include all retina visible at a magnification of 100×, 2. Full retinal thickness defined as extending from the internal limiting membrane to the apex of the retinal pigment epithelium, or 3. Ten thickness measurements taken in the defined field to determine mean retinal thickness. Consistent application of these limitations will assure reliable and reproducible results. Similar approaches can be applied to any organ. *Substantial portions of the text in this section are taken from Smith, R.S., Martin, G., and Boggess, D., Kinetics and morphometrics, in Systematic Approach to Evaluation of Mouse Mutations, Sundberg, J.P. and Boggess, D., Eds., CRC Press, Boca Raton, FL, 2000, chap. 8.

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Quantitative Morphometrics Phenotypic differences between mouse strains are often initially observed as variations in morphological features. Distinguishing putative interstrain differences from normal variation between individuals of the same strain requires some sort of quantification of the anatomical differences. Characterizing the mutant phenotype may also involve quantifying relevant morphological features. Relevant features may be at any level of organization: gross, organ, tissue, cell, or subcellular. Morphometric data can be collected in several different ways, depending on the nature of the relevant morphological features under consideration as well as the degree of variation observed. Obvious differences in some readily measurable morphological feature may permit very straightforward analysis. Examples would be large differences in length of structures or thickness of tissue layers visualized in tissue sections. Morphometric data can be collected in these instances using a measuring scale in the ocular of the microscope or with a ruler and photographic images. Standard statistical analysis can be used to determine averages, variation, and significant differences. It may, however, be necessary to measure more complex feature parameters such as area, perimeter, number, or lengths of curved objects. In addition, it may often be more meaningful to express these parameters in relation to the real world of three dimensions. Thus, areas become volumes, and perimeters become surface areas. This information, as well as one-dimensional data such as number and length, is best expressed in relation to total area or volume and requires more advanced techniques. One of the most powerful methods to achieve these ends is stereology. Stereology is a method for obtaining quantitative 3D morphological data from two-dimensional images of sectioned material. Although a discussion of the principles and techniques of stereology is beyond the scope of this discussion, several reviews are available.22-24 The technique provides the ability to quantify tissue elements in terms of their number, length, surface area, or volume in relation to some reference parameter, typically the total volume of the tissue type under investigation. The basic principle of stereology is the use of standardized test grids applied to images of sectioned material. Counting the number of interactions between the test grid and the features of interest provides data that can be used in well-established equations to derive three-dimensional quantitative information in an efficient manner. The use of complex specimen parameters such as color and density to describe and quantify morphological features requires the use of computerized image analysis. In this case, methods of specimen preparation must provide images in which the structures of interest can be readily differentiated from the surrounding tissue by computer-based image analysis techniques. Such techniques may include fluorescence- or colorimetric-based immunolabeling of particular tissue elements. Standard or specialized histochemical stains may be used to dye selectively particular cell types or tissue elements. These specimens can then be analyzed using digital imaging and computer-aided segmentation. In all cases it is essential that the computer programs have facilities for operator interaction, both in terms of defining the means by which the computer selects tissue elements for analysis (such as color, intensity, or shape) and of providing for the ability to edit the data set to correct for errors both in excluding the structures of interest and in including irrelevant structures. The use of computerized image analysis to aid in obtaining morphometric data promises to be valuable in more straightforward analysis as well. Digital imaging produces images that are immediately available for analysis on the computer screen. These images can be calibrated to real dimensions and data can be obtained quite efficiently, using the computer mouse as a “digital ruler” to define the boundaries of the tissue elements to be measured. These data can be readily tabulated for analysis by spreadsheet or statistical software. More complex measurements such as areas or proportional area can also be generated for either used-defined or computer-segmented regions. Studies indicate that the combined use of confocal microscopy, 3D reconstruction, and computer-assisted morphometric analysis of thick sections25 and living tissue26 provides information

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comparable to that obtained by standard histological preparations. Used singly or in combination, these are extremely powerful tools for investigating the crucial interrelationships of structure and function in the eye that often must be examined simultaneously with multiple indicators and in multiple dimensions.

CELL KINETICS* Determination of relative rates of cell proliferation is often important in evaluating mouse mutations. For example, when phenotypic differences are subtle, it may become necessary to quantitate cellular activity to differentiate between homozygotes, heterozygotes, and wild-type mice. The ability to determine significant quantitative differences is particularly important with phenotypes that are influenced by modifier genes (quantitative traits). These measurements are also useful in deciding whether a thickened cell layer is due to enhanced proliferation or to decreased cell death.27 Measurements may require simple counts of labeled cells or more complex morphometric analyses. Several techniques can be used to measure DNA synthesis, mitotic indices, and cell cycle rates. Single or multiple injections of a substance that can be detected in the cell nucleus is called pulse labeling. Other immunohistochemical techniques can be used to measure DNA synthesis in archival paraffin-embedded sections. Actively growing cells pass through the cell cycle, which has four main steps: G1, the relatively inactive period following mitosis; S, the stage of DNA synthesis; G2, the postsynthetic/premitotic phase; and M, the mitotic phase. Pulse-labeling methods indicate the number of cells in the DNA synthetic phase (Figure 14A.8).28

Cyclin A&B

3 HT BrdU PCNA*

S

*methanol fix

G2 Ki67 PCNA (non-methanol fix) (All cycling cells)

G1

M

Mitotic figure counts

Cyclin D&F

FIGURE 14A.8 Cell cycle.

Ki-67 Antigen This monoclonal antibody detects a nonhistone protein nuclear antigen present in proliferating cells but absent in quiescent cells.39 Ki-67 is visualized using standard avidin–biotin–peroxidase techniques from commercially available kits. It has been used for assessing prognosis of some tumors and may be a more specific indicator of cell proliferation than PCNA, even though the labeling is not restricted to S-phase cells.40 The number of labeled cells correlates well with tritiated thymidine labeling.35 However, we have found that fewer nuclei are stained with Ki-67 than with the techniques described below. As an example (Figure 14A.9A), the lens epithelium demonstrates prominent stain*Substantial portions of the text in this section are taken from Smith, R.S., Martin, G., and Boggess, D., Kinetics and morphometrics, in Systematic Approach to Evaluation of Mouse Mutations, Sundberg, J.P. and Boggess, D., Eds., CRC Press, Boca Raton, FL, 2000, chap. 8.

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ing, but proliferating cells of the epidermis failed to stain, although positive results were obtained with tritiated thymidine, BrdU, and PCNA in sections from the same tissue. Proliferating Cell Nuclear Antigen (PCNA) PCNA is an acidic nuclear protein that is synthesized prior to mitosis.33 No treatment of the experimental animal is required for demonstration of PCNA expression. Cells fixed in either 4% formaldehyde in 0.1 M cacodylate buffer or in absolute methanol demonstrate the most-pronounced staining (Figure 14A.9B).34 The method of fixation is important, since PCNA labeling with methanol fixation is limited to the S-phase cells, whereas other fixatives produce staining of any cycling cell and are less specific.35 Questions have been raised concerning the validity of PCNA staining when compared to BrdU staining for assessing tumor-related mortality.36 Despite these objections, PCNA staining offers the unique advantage that archival paraffin-embedded tissues can be used to identify cycling cells, even in tissues that were not fixed in methanol.37,38 Bromodeoxyuridine Bromodeoxyuridine (BrdU) is a nonradioactive alternative for labeling cells entering the S-phase. Mutant and control mice are injected intraperitoneally with 50 µg/g body weight BrdU. After 1 h, mice older than 12 days are euthanized by CO2 asphyxiation and tissues are removed and placed in fixative for 12 to 24 h. Mice up to 7 days of age are held for 2 h before tissue collection, since the extra time enhances the intensity of the BrdU staining. Standard paraffin sections are cut at 5 mm and stained with hematoxylin and eosin for morphologic evaluation. Sequential unstained sections are placed on slides coated with poly-L-lysine to minimize folding and tissue loss and stained with a monoclonal anti-BrdU antibody. The positive staining nuclei (dark brown nuclei) are counted (Figure 14A.9C). Labeling of cells with either BrdU or 3H-thymidine produces comparable results.32

FIGURE 14A.9 A. Normal mouse lens epithelium stained with Ki67 antibody. Only seven cells are stained. B. Normal mouse lens epithelium stained with PCNA; nearly all cells are positive. C. Normal mouse lens epithelium stained with BrdU; half of the cells are stained. D. Normal mouse lens epithelium after tritiated thymidine exposure; nearly half the cells demonstrate uptake of the radiolabel. Arrows indicate selected positive cells. Original magnification × 630.

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Radioactive Labeling This technique has been used successfully for over 50 years. Thymidine is incorporated into DNA during the S-phase of the cell cycle and can be isotopically labeled with tritium (3H) or carbon (14C). Usually, a single dose is administered intraperitoneally, followed by tissue collection after 1 to 2 h. Depending on experimental design, multiple pulses of 3H-thymidine may be required, a technique that has been used to identify the location of conjunctival and cutaneous stem cells.29,30 The usual dosage for mice is 1 to 5 µCi of 3H-thymidine per gram body weight.27,29,31 Any cell in the S-phase exposed to the 3H-thymidine will incorporate the isotope. Standard precautions and techniques for working with radioactive materials should be observed when using these techniques. After collection and routine fixation and embedding of the eyes, paraffin sections are prepared and covered with a liquid photographic emulsion under darkroom conditions (Kodak NTB-2, Eastman Kodak Co., Rochester, NY). The slides are routinely exposed for 30 days at 4°C (to optimize visualization, extra slides can be cut and exposed for 25, 30, 35, and 40 days), developed (Kodak D19 developer, Eastman Kodak Co.), and counterstained with hematoxylin.30 Cells with more than four grains per nucleus are considered positive and these cells are counted and expressed as positive cells per unit area (Figure 14A.9D). The disadvantages of this technique include (1) the training required to work with radioactive materials, (2) potential biohazards, and (3) the wait of 4 to 6 weeks for exposure of the emulsion. In addition, there is always diffuse background tissue labeling that may be confusing in determining positive vs. negative cell labeling, although the visualization is enhanced by the use of dark-field microscopy. Mitotic Rates Mitotic activity can be measured and reported as the number of mitotic figures per high-power field or for a defined area. This complements the previously described methods, since it measures a different part of the cell cycle (mitotic phase). Although many histochemical stains are available, hematoxylin alone provides the best contrast for evaluating and counting mitotic figures. General Problems of Interpretation In both tritiated thymidine and BrdU labeling, a comparison is usually made between tissues from normal controls and mutant mice. Care must be taken that the “controls” correctly fit the term. An example of an inappropriate control would be a mouse heterozygous for an allele that produces a heterozygote effect: an animal that is phenotypically normal by clinical examination may still have microscopic phenotypic differences from a true wild-type mouse. Location of tissue sampled must be held constant between control and affected mice. A sample of dorsal skin cannot be compared with a sample of ventral skin. In many organs, such as the eye, tissue orientation and plane of section also need to be constant. For example, if the corneal epithelium is measured, the results will be different if the section used contains central or peripheral cornea.27 An important part of the experimental protocol is to define what is being measured, where it is being measured, and how it is being measured. If these parameters are not strictly controlled, conclusions will be meaningless Since many labeling techniques mark cells in the S-phase of growth, these methods are often used to provide quantitative measurements of cell proliferation. The process of mitosis is also easily detected, enabling calculation of the mitotic index. In both instances, the first step is to define the area of measurement (e.g., labeled cells/linear mm of tissue; labeled cells touching the basement membrane; labeled cells per 400× field, etc.) A similar approach would be to count the number of labeled or mitotic cells/1000 total cells counted.41 Any of these procedures provides the number of marked cells per unit of measurement, which allows statistical analysis of experimental results. It should be emphasized that well-defined parameters of measurement must be established and used with care, if useful quantitative data are the goal.42

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REFERENCES 1. Minsky, M., Memoir on inventing the confocal scanning microscope, Scanning, 10:128, 1988. 2. Sheppard, C.J.R. and Shotton, D.M., Confocal Laser Scanning Microscopy, BIOS Scientific Publishers, Oxford, 1997. 3. Pawley, J.B., Ed., Handbook of Biological Confocal Microscopy, 2nd ed., Plenum Press, New York, 1995. 4. Wilson, T. and Sheppard, C.J.R., Theory and Practice of Scanning Optical Microscopy, Academic Press, London, 1984. 5. Brotchie, D. et al., Characterization of ocular cellular and extracellular structures using confocal microscopy and computerized three-dimensional reconstruction, Meth. Enzymol., 307:496, 1999. 6. Mason, W.T., Ed., Fluorescent and Luminescent Probes for Biological Activity, 2nd ed., Academic Press, New York, 1999. 7. Tatton, N.A. et al., A fluorescent double-labeling method to detect and confirm apoptotic nuclei in Parkinson’s disease, Ann. Neurol., 44:S142, 1998. 8. Stevens, J.K. et al., Eds., Three-Dimensional Confocal Microscopy: Volume Investigation of Biological Systems, Academic Press, New York, 1994. 9. http://www.cs.ubc.ca/spider/ladic/confocal.html. 10. D’Amato, R., Wesoloski, E., and Hodgson Smith, L.E., Microscopic visualization of the retina by angiography with high-molecular-weight fluorescein-labeled dextrans in the mouse, Microvasc. Res., 46:135, 1993. 11. Cody, S.H. and Williams, D.A., Optimizing confocal microscopy for thick biological specimens, in Fluorescent and Luminescent Probes for Biological Activity, 2nd ed., Mason, W.T., Ed., Academic Press, New York, 1999. 12. Furrer, P., Mayer, J.M., and Gurny, R., Confocal microscopy as a tool for the investigation of the anterior part of the eye, J. Ocul. Pharmacol. Ther., 13:559, 1997. 13. Maurer, J.K. and Jester, J.V., Use of in vivo confocal microscopy to understand the pathology of accidental ocular irritation, Toxicol. Pathol., 27:44, 1999. 14. Bohnke, M. and Masters, B.R., Confocal microscopy of the cornea, Prog. Retin. Eye Res., 18:553, 1999. 15. Fitzke, F.W., Imaging the optic nerve and ganglion cell layer, Eye, 14:450, 2000. 16. Yaylali, V. et. al., In vivo confocal imaging of corneal neovascularization, Cornea, 17:646, 1998. 17. Jester, J.V. et al., Inhibition of corneal fibrosis by topical application of blocking antibodies to TGF beta in the rabbit, Cornea, 16:177, 1997. 18. Moritz, O.L. et al., Fluorescent photoreceptors of transgenic Xenopus laevis imaged in vivo by two microscopy techniques, Invest. Opthamol. Vis. Sci., 40:3276, 1999. 19. Haugland, R.P. and Johnson, I.D., Intracellular ion indicators, in Fluorescent and Luminescent Probes for Biological Activity, 2nd ed., Mason, W.T., Ed., Academic Press, New York, 1999. 20. Hirata, K., Nathanson, M.H., and Sears, M.L., Novel paracrine signaling mechanism in the ocular ciliary epithelium, Proc. Natl. Acad. Sci., U.S.A., 95:8381, 1998. 21. Bazan, H.E. et. al., Platelet-activating factor induces cyclooxygenase-2 gene expression in corneal epithelium. Requirement of calcium in the signal transduction pathway, Invest. Opthamol. Vis. Sci., 38:2492, 1997. 22. Russ, J.C. and Dehoff, R.T., Practical Stereology, 2nd ed., Plenum Press, New York, 2000. 23. Bolender, R.P., Hyde, D.M., and Dehoff, R.T., Lung morphometry: a new generation of tools and experiments for organ, tissue, call, and molecular biology, Am. J. Physiol., 265:L521, 1993. 24. Mayhew, T.M., A review of recent advances in stereology for quantifying neural structure, J. Neurocytol., 21:313, 1992. 25. Camp J.J. et al., Three-dimensional reconstruction of aqueous channels in human trabecular meshwork using light microscopy and confocal microscopy, Scanning, 19:258, 1997. 26. Pater, S.V. et. al., automated quantification of keratocyte density by using confocal microscopy in vivo, Invest. Opthalmol. Vis. Sci., 40:320, 1999. 27. Smith, R.S. et al., Corn1: a mouse model for corneal surface disease and neovascularization, Invest. Ophthalmol. Vis. Sci., 37:397, 1996.

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28. Scragg, M.A. and Johnson, N.W., Epithelial cell kinetics, J. Oral Pathol., 11:102, 1982. 29. Wei, Z. et al., Label-retaining cells are preferentially located in fornical epithelium: Implications on conjunctival epithelial homeostasis, Invest. Ophthalmol. Vis. Sci., 36:236, 1995. 30. Cotsarelis, G., Sun, T.T., and Lavker, R.M., Label-retaining cells reside in the bulge area of pilosebaceous unit: implications for follicular stem cells, hair cycle, and skin carcinogenesis, Cell, 61:1329, 1990. 31. Sundberg, J.P. et al., Full-thickness skin grafts from flaky skin mice to nude mice: maintenance of the psoriasiform phenotype, J. Invest. Dermatol., 102:781, 1994. 32. del Rio, J.A. and Soriano, E., Immunocytochemical detection of 5′-bromodeoxyuridine incorporation in the central nervous system of the mouse, Exp. Brain Res., 49:311, 1989. 33. Gao, C.Y. and Zelenka, P., Cyclins, cyclin-dependent kinases and differentiation, Bioessays, 19:307, 1997. 34. Galand, P. and Degraef, C., Cyclin/PCNA immunostaining as an alternative to tritiated thymidine pulse labeling for marking S-phase cells in paraffin sections from animal and human tissues, Cell Tissue Kinet., 22:383, 1989. 35. Hofstadter, F., Knuchek, R., and Ruschoff, J., Cell proliferation assessment in oncology, Virchows Arch., 427:323, 1995. 36. Ghazvini, S. et al., Comparative analysis of proliferating cell nuclear antigen, bromodeoxyuridine, and mitotic index in uveal melanoma, Invest. Ophthalmol. Vis. Sci., 36:2762, 1995. 37. Greenwell, A., Foley, J.F., and Maronpot, R.R., An enhancement method for immunohistochemical staining of proliferating cell nuclear antigen in archival rodent tissues, Cancer Lett., 59:251, 1991. 38. Greenwell, A., Foley, J.F., and Maronpot, R.R., Detecting proliferating cell nuclear antigen in archival rodent tissues, Environ. Health Perspect., 101(Suppl. 5):207, 1991. 39. Gerdes, J. et al., Immunobiochemical and molecular biological characterization of the cell proliferationassociated nuclear antigen that is defined by the monoclonal antibody Ki-67, Am. J. Pathol., 138:867, 1991. 40. Abele, M.C. et al., Significance of cell proliferation index in assessing histological prognostic categories in Hodgkin’s disease, Haematologica, 82:281, 1997. 41. Plumb, J.A. and Wright, N.A., Epidermal cell population kinetics, in Methods in Skin Research, Skerrow, D. and Skerrow, C.J., Eds., John Wiley & Sons, New York, 1985, chap. 10. 42. Sundberg, J.P. et al., Harlequin icthyosis (ichq): a juvenile lethal mouse mutation with icthyosiform dermatitis, Am. J. Pathol., 151:293, 1997.

B. INTRAOCULAR PRESSURE MEASUREMENT IN MICE: TECHNICAL ASPECTS Simon W. M. John and Olga V. Savinova

INTRODUCTION Mice are expected to be extremely helpful in characterizing genes and mechanisms that affect intraocular pressure (IOP).1 After a brief introduction to mouse IOP, this section focuses on technical aspects of IOP assessment in mice and does not detail published findings about IOP values and genes or other factors that alter IOP. The aim is to address various technical questions that we have been asked over the past few years. Based on over 35 mouse strains studied, average IOP in nonglaucomatous mouse strains ranges from approximately 10 to 20 mmHg (Figure 14B.1 and References 2 and 3). This range is similar to that in human populations. Gender does not typically affect IOP in normal strains and most tested strains exhibit a diurnal rhythm with IOP being the highest during the dark period of the day.2 Some strains develop elevated IOP and glaucoma with age.4,5

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22 20

Females

Males & Females

Males

C57BLKS/J DBA/1J CBA/CaHN


SB/Le C57BL/6J ST/bJ SM/J C57BL/10J A/HeJ SWR/J SF/CamEi CE/J LP/J

314

18 16 14 12 10 8 6 4 2 0

RIIIS/J SEC/IReJ BALB/cJ BALB/cByJ SJL/J NZW/LacJ RF/J MA/MyJ C58/J C3H/HeJ 129P3/J CAST/Ei A/J C57L/J C57BL/6ByJ BUB/BnJ I/LnJ NZB/B1NJ AKR/J CBA/J CBA/CaJ

(mmHg)

Intraocular Pressure

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Strain FIGURE 14B.1 IOP differences between genetically distinct mouse strains. Mean IOP ± SEM is shown for each strain. For each strain, except for RIIIS/J (7 mice), 12 to 25 mice were analyzed. Mice of most strains were 6 to 12 months old, except for SEC/1ReJ and SB/Le that were 2 to 3 months old. CBA/CaHN-Btkxid/J is abbreviated to CBA/CaHN. (Courtesy of BioMed Central.2)

IMPORTANT DIMENSIONS AND VOLUMES Although there is some variation with strain and age, the diameter of mouse eyes is approximately 3 mm and the normal corneal radius of curvature (external) is approximately 2.8 mm2 (assessed by fitting specially fabricated contact lenses; R. Smith, personal communication). Ultrasound analysis of live mice shows that the normal anterior chamber is around 350 µm deep at its deepest location4 and the cornea is approximately 75 to 100 µm thick. We have measured the volume of aqueous humor to be 5.8 ± 0.3 µl in strain C57BL/6J mice (n = 12 eyes ) and 5.1 ± 0.4 µl in strain C3HeB/FeJ mice (n = 9 eyes). Because of these small dimensions, we initially thought that a servo-nullifying method designed to measure pressure in very small volume compartments might be required to measure mouse IOP.6-9 Initial attempts to penetrate the cornea with the very fine microcannulas required for this method (5 µm or smaller diameter) were unsuccessful and making the fine electrodes was not straightforward. Thus, we developed and evaluated a manometric technique involving direct measurement of pressure following cannulation of the anterior chamber.2,10 The cannula is a microneedle that has an overall diameter (OD) of approximately 50 µm that is inserted 50 to 100 µm into the anterior chamber. The volume occupied by the inserted, fluid-filled needle tip is approximately 0.0002 µl if the needle is inserted 100 µm into the eye (volume occupied = πr2h µm3 or πr2h × 10-9 µl; for a 50-µm OD microneedle = 3.142 × 252 × 100 × 10-9 = 0.0002 µl; r = radius, h = length inserted). Although great care needs to be taken to make accurate measurements, this procedure is reliable and produces reproducible data over extended periods of time.2 It is accurate and rapid enough to allow large-scale genetic studies of factors determining IOP.2

MEASUREMENT PROCEDURE The measurement procedure, equipment, and microneedle fabrication have been described in detail.10 A schematic representation of the system is shown in Figure 14B.2, and the actual instrumentation is shown in Figure 14B.3. The mouse is anesthetized and placed on the platform and gently held in

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Pressure Transducer

Reservoir Valve

Amplifier

Analogue to Digital Translater

Computer Interface

Computer

FIGURE 14B.2 Diagrammatic representation of the equipment used to measure IOP. A modification compared to our original arrangement10 is that the microneedle is directly attached to the transducer without tubing.

position with stainless steel holders. Care is taken to avoid pressure on the neck that could alter IOP. A drop of isotonic saline is placed on each eye. The saline prevents corneal dehydration and alters refraction to allow a clear view into the anterior chamber. The left eye is then viewed under a microscope and the microneedle tip is placed inside the drop of saline and the pressure reading is zeroed. (Right eye measurements are possible but we have not routinely assessed the right eye.) The tip of the microneedle is then inserted into the anterior chamber by piercing the cornea over the pupil. To facilitate penetration, the eye is gently stabilized using a curved wire instrument that cradles the eye and has a handheld handle. Although position may vary with investigator, the needle is inserted into the eye from the nasal direction. It is inserted at an angle of 40 to 50° from a vertical, central axis running from anterior to posterior through the eye, and an angle of approximately 40° horizontal, above a central axis running from cranial to caudal sides of the eye. The stabilization instrument is placed against the eye wall in a position opposing the microneedle during corneal penetration, and with developed skill the microneedle easily enters the eye. Exact entry position varies from mouse to mouse, and is determined on a case-by-case basis by watching how the eye and microneedle move to achieve a smooth entry. Ocular entry takes skill and is the most difficult part of the procedure to master. Care is taken to minimize corneal deformation and to ensure that the eye remains in its normal position. While establishing the procedure it was important to determine if IOP is altered upon entering the eye. To do this a duplicate measurement system was constructed (Figure 14B.4). IOP measurement was performed as usual, except that the microneedle was left in the anterior chamber after obtaining a pressure reading (IOP1). The microneedle of the second system was then inserted into the anterior chamber. The pressure recorded via the second microneedle (IOP2) was then compared to the reading obtained prior to its insertion. This analysis was performed in 15 different eyes (Figure 14B.4) and showed that pressure was not substantially altered upon entering eyes with pressures in the normal range (mean –0.3 ± 0.3 mmHg).10 At the end of the measurement, gentle pressure is applied to the eyelid to confirm microneedle patency. A prompt increase in pressure is required for inclusion of data. Visual inspection of the microneedle and prompt zeroing on removal from the eye are also required. In cases where penetration of the cornea is difficult or more than gentle pressure is applied to the eye the data are discarded, as the IOP may have been altered. For inclusion of data, it is also required that an eye has minimal or no leakage. An eye is regarded as leaky if IOP readings drop rapidly or more than 1 mmHg over the

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FIGURE 14B.3 Instrumentation used to measure IOP. A. A photograph demonstrating the relative positions of the various pieces of equipment. The platform on which the mouse is placed is at a slight angle that is always kept constant. B. A close-up view. The head stabilizer (1) is made from steel wire that is silver-soldered and is attached to one of the light sources. The snout stabilizer (2), the body holder (3) which prevents the mouse from slipping, and the needle holder (4) are labeled. The needle holder is the World Precision Instruments MPH6S electrode holder that is modified by placing a second gasket in the luer-accepting end. The needle is threaded through both gaskets, and the space between the gaskets that is outside of the needle is filled with water to prevent air from entering the system. C. The mouse rests on the platform and the snout and head stabilizers are used to help position the head, avoiding pressure on the neck. The eye is covered with a drop of saline.

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measurement period. Most eyes do not leak substantially during the measurement period (Figure 14B.4) and we typically succeed in measuring IOP in over 80% of the eyes in an experiment. Prolonged measurements are not often possible as leakage starts after a few minutes, as a result of ocular movement against the needle as the mouse breathes. This problem may be preventable by allowing the needle to move with the eye but we have not yet tested this. On removal of the needle, many eyes leak and so it is not possible to make further measurements in a short time period. The eyes recover quickly with no inflammation or long-term detrimental effects. Multiple measurements in the same eye are possible but we have not extensively tested this. We have measured the IOP as many as three times in the same groups of mice at intervals of 2 to 3 months with comparable results. For example, IOP differences between groups of mice that are homozygous mutant (c-2J allele) or homozygous normal at the tyrosinase locus (Tyr)2 were similar when IOP was measured for the first time (Tyrc-2J/c-2J 14.2 ± 0.4 mmHg, Tyr+/+ 12.4 ± 0.3 mmHg, P < 0.001) or second time (Tyrc-2J/c-2J 14.8 ± 0.5 mmHg, Tyr+/+ 12.9 ± 0.3 mmHg, P = 0.001).

A

etc.

IOP2

IOP1 Amplifier

Analogue to Digital Translater

B

Computer Interface

Computer

C 8 6

10

IOP change after microneedle insertion

4 2 0 -2 -4 -6 -8 -10

IOP change (mmHg)

10

IOP change (mmHg)

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IOP change over measurement period

4 2 0 -2 -4 -6 -8 -10

Observations

Observations

FIGURE 14B.4 System checks. A. Duplicate measurement systems were used to assess the effects of microneedle insertion on IOP. The pressure recorded with the first microneedle inserted (IOP1) was compared to the pressure recorded through the second microneedle (IOP2) and the results are shown in B. There was minimal effect of needle insertion in these already cannulated eyes that may be more prone to leakage than cannulated eyes. C. The change in pressure recorded during a 90-s measurement period for 20 representative mice of four different strains is shown.

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ANESTHESIA Many anesthetic agents, including xylazine, lower IOP. Ketamine usually appears to increase IOP,11-13 but there are reports of ketamine having no effect on IOP or even reducing IOP.11,14 Different doses, routes of administration, or environments may contribute to these differences. Initially, we attempted to control for potential effects of anesthesia by administering anesthetic to one mouse at a time and carefully monitoring the plane of anesthesia.10 Because the relationship between measured IOPs and those in conscious mice depends upon the effect of the anesthetic protocol, subsequently we have investigated the effect of our anesthesia protocol on IOP (intraperitoneal injection of 99 mg/kg ketamine and 9 mg/kg xylazine). Despite a depressant effect on IOP by 25 min, our anesthetic protocol has no detectable effect on IOP during the first 12 min after administration (Figure 14B.5 and Reference 2). Thus, to avoid effects of anesthesia on IOP, all measurements are made within a window of up to 12 min after anesthetic administration. It is worth mentioning that environment can alter the effect of anesthesia. Cage cleanliness, changing frequency, and housing density can alter drug metabolism and the effect of anesthesia in rats.15,16 The type of bedding used is also important. We use wood shavings for bedding and wood shavings expose the mice to terpenes. Terpene administration or environmental exposure to terpenes in wood shavings alters drug resistance and decreases the effect of anesthetic agents in both rats and mice.17-22

Intraocular Pressure (mmHg)

16 14

A

12 10 8 6 4

Dose 1.0 x 1.5 x 2.0 x

2 0 25

5 18

B

16

Intraocular Pressure (mmHg)

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IOP = 13.354 - 0.006 x Time; R2 = 1.5 x 10-4 0 4

5

6

7

8

9

Time (min)

10

11

12

FIGURE 14B.5 Effects of anesthesia. A. Mean IOP ± SD is shown for C57BL/6J mice at 5 and 25 min after administration of various doses of anesthetic. The 1× dose consisted of 99 mg/kg ketamine and 9 mg/kg xylazine. All doses decreased IOP by 25 min. At all doses the IOP at 5 min was the same suggesting that the effect of anesthesia had not yet occurred. Approximately 30 3- to 4-monthold mice were analyzed at each dose and time. B. Scatterplots demonstrating no change of IOP in 195 C57BL/6J mice during the 12 min following anesthetic administration. The mice were 3 to 6 months old and the sexes and ages were equally represented at each time point. Similarly, no change of IOP has been noted during this period in other tested strains.2 (Courtesy of BioMed Central.2)

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STRESS AND OTHER FACTORS In general, the mean IOP of any specific strain has been highly reproducible over time, typically differing by only 1 to 1.5 mmHg.2 Although not specifically tested, we have noticed that stress can increase IOP. For example, the mean IOP of C57BL/6J mice is typically around 13 mmHg but, in cages housing males that were observed to fight, the mean IOP is around 17 mmHg. Similarly, increased IOP was noticed during periods of construction close to our animal facility. To avoid effects of stress and environment, we make our measurements in a sound-insulated procedure room and the mice are typically housed in the room for 1 to 2 weeks before measurement so that they are acclimatized to the environment. Obviously, diet should be controlled as the effects of various diets or dietary components on IOP are not known. Most strains we have tested demonstrate diurnal variation of IOP,2 and so it is important to assess groups to be compared at the same time of day. When planning experiments, strain-specific characteristics such as susceptibility to disease should be considered as they may affect the data. Strain 129P3/J (formerly 129/J) is the one of the most variable nonglaucomatous strains we have assessed with average IOP ranging from approximately 14 to 16.5 mmHg, with some mice having IOPs greater than 20 mmHg. This strain is susceptible to blepharoconjunctivitis23 which tends to occur at an earlier age than in other susceptible strains (S. John, personal observation). Thus, it is possible that unnoticed conjunctival inflammation sometimes results in increased IOP and contributes to the variability of IOP in this strain.

REFERENCES 1. John, S.W.M., Anderson, M.G., and Smith, R.S., Mouse genetics: a tool to help unlock the mechanisms of glaucoma, J. Glaucoma, 8:400, 1999. 2. Savinova, O.V. et al., Intraocular pressure in genetically distinct mice:an update and strain survey, BMC Genet., http://www.biomedcentral.com/1471-2156/2/12/. 3. Smith, R.S. et al., Haploinsufficiency of the transcription factors FOXC1 and FOXC2 results in aberrant ocular development, Hum. Mol. Genet., 9:1021, 2000. 4. John, S.W.M. et al., Essential iris atrophy, pigment dispersion, and glaucoma in DBA/2J mice, Invest. Ophthalmol. Vis. Sci., 39:951, 1998. 5. Anderson, M.G. et al., Genetic modification of glaucoma associated phenotypes between AKXD-28/Ty and DBA/2J mice, BMC Genet., 2, 2001. Available at www.biomedcentral.com/1471-2156/2/1/. 6. Mäepea, O. and Bill, A., Pressures in the juxtacanalicular tissue and Schlemm’s canal in monkeys, Exp. Eye Res., 54:879, 1992. 7. Mäepea, O. and Bill, A., The pressures in the episcleral veins, Schlemm’s canal and the trabecular meshwork in monkeys: effects of changes in intraocular pressure, Exp. Eye Res., 49:645, 1989. 8. Fein, H., Microdimensional pressure measurements in electrolytes, J. Appl. Physiol., 32:560, 1972. 9. Wiederhielm, C.A. et al., Pulsatile pressures in the microcirculation of frog’s mesentery, Am. J. Physiol., 207:173, 1964. 10. John, S.W.M. et al., Intraocular pressure in inbred mouse strains, Invest. Ophthalmol. Vis. Sci., 38:249, 1997. 11. Antal, M., Mucsi, G., and Faludi, A., Ketamine anesthesia and intraocular pressure, Ann. Ophthalmol., 10:1281, 1978. 12. Marynen, L. and Libert, Ocular tonometry in the child under general anesthesia with IM ketamine, Acta Anaesthesiol. Belg., 27:29, 1976. 13. Bar Ilan, A. and Pessah, N.I., On the use of ketamine in ocular pharmacological studies, J. Ocul. Pharmacol., 2:335, 1986. 14. Erickson Lamy, K.A. et al., Comparative anesthetic effects on aqueous humor dynamics in the cynomolgus monkey, Arch. Ophthalmol., 102:1815, 1984. 15. Vessell, E.S. et al., Hepatic drug metabolism in rats: impairment in a dirty environment, Science, 179:896, 1973. 16. Einon, D. et al., Effect of isolation on barbiturate anaesthesia in the rat, Psychopharmacology (Berlin), 50:85, 1976.

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17. Pick, J.R. and Little, J.M., Effect of type of bedding material on thresholds of pentylenetetrazol convulsions in mice, Lab. Animal Care, 15:29, 1965. 18. Vesell, E.S., Induction of drug-metabolizing enzymes in liver microsomes of mice and rats by softwood bedding, Science, 157:1057, 1967. 19. Brochet, D. et al., Effects of single intraperitoneal injections of an extract of Ginko biloba (EGb 761) and its terpene trilactone constituents on barbital-induced narcosis in the mouse, Gen. Pharmacol., 33:249, 1999. 20. Malkinson, A.M., Prevention of butylated hydroxytoluene-induced lung damage in mice by cedar terpene administration, Toxicol. Appl. Pharmacol., 49:551, 1979. 21. Malkinson, A.M. and Shere, W.C., Decreased pentabarbital sleeptime following a single intraperitoneal injection of cedar-derived sesquiterpenes, Res. Commun. Chem. Pharmacol., 25:607, 1979. 22. Wade, A.E. et al., Alteration of drug metabolism in rats and mice by an environment of cedarwood, Pharmacology, 1:317, 1968. 23. Smith, R.S., Montagutelli, X., and Sundberg, J.P., Ulcerative blepharitis in aging inbred mice, in Pathobiology of the Aging Mouse, Mohr, U. et al., Eds., ILSI Press, Washington, D.C., 1996, 131.

C. ELECTROPHYSIOLOGICAL TESTING OF THE MOUSE VISUAL SYSTEM Steven Nusinowitz, William H. Ridder III, and John R. Heckenlively

INTRODUCTION The abundance of new and different inbred strains, knockouts, and transgenic mouse models of human ocular disease has encouraged the development of objective, reliable, and rapid methods of assessing the functional status of the visual system in these mice. This chapter reviews several electrophysiological techniques for assessing the functional status of the retina and visual system in the mouse, including the full-field electroretinogram (ERG), the multifocal electroretinogram (mERG), and visually evoked cortical potentials (VECP). The Full-Field Electroretinogram The ERG is an electrical signal that is easily recorded from the corneal surface of the mouse eye and represents the massed response of the retina to light stimulation. The solid curve shown in Figure 14C.1 is an example of an ERG recording from a normal C57BL/6J mouse in response to a bright flash. The major components of the response are the a-wave, which is the first negative corneal potential, and the b-wave, which is the first positive corneal potential. ERGs to bright flashes presented in the dark also contain a high-frequency oscillatory component on the ascending limb of the b-wave, collectively called the oscillatory potentials (OPs). After the onset of steady illumination, the relatively fast a- and b-waves are followed by a slower, positive-going cwave (not shown in Figure 14C.1). Considerable effort has been devoted to understanding the cellular origin of the different components of the ERG. Under dark-adapted conditions, the leading edge of the a-wave is generally associated with rod photoreceptor activity.1-3 The b-wave is associated with the combined activity of depolarizing bipolar cells and bipolar cell-dependent K+ currents affecting Muller cells.4-9 The cellular origins of the OPs are not completely understood, although they are likely generated by amacrine cells and other inner retinal cells interacting with bipolar and ganglion cells.10-14 The c-wave of the ERG is a corneal positive potential recorded across the retinal pigment epithelium (RPE) and results from an increase in the transepithelial potential (TEP) of the RPE.15 Because of the technical difficulties in recording the c-wave, it has not found general use in the mouse. Contributions from amacrine and ganglion cells have also been identified in the scotopic threshold response (STR), which is a negative-going potential in the dark-adapted ERG that is present at threshold and with dim illumination.16 The STR has been recorded from human, primate, cat, and rat

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FIGURE 14C.1 Representative ERG response to a bright flash obtained from a normal adult C57BL/6J mouse (see text for details).

retinas, but so far there are no publications demonstrating such recordings from the mouse (for rats, see References 17 and 18). Basic ERG Recording Technique In laboratories where mouse ERGs are currently recorded, different recording techniques (e.g., electrodes), methods of stimulating the eye (e.g., Ganzfeld vs. Maxwellian view), and experimental protocols are employed.19-24 Because a standard protocol for mice similar to that adopted for humans is currently unavailable, the differences in methodology can make comparisons of results between laboratories and animal models difficult. In principle, the techniques for recording the ERG from the mouse are virtually identical to those used in human studies. The major components of a typical ERG system are (1) a light source for stimulating the retina, (2) electrodes for recording the signal generated by the retina in response to light, (3) a signal amplification system, and (4) a data acquisition system to accumulate, condition, and display data. Most commercial systems have integrated all of these elements into a single unit. However, it is a relatively easy task to assemble a data acquisition system from individual components provided that one possesses a moderate level of electronics and computer programming skills. While commercial systems are ready made and easy to use, their major disadvantages are that they are expensive, they are designed primarily for human clinical applications, and they lack the flexibility for adjusting conditions and testing protocols. The major components of the ERG system used in our laboratory are shown in Figure 14C.2. The mouse eye is positioned at the opening of a spherical dome (A) whose interior surface is painted with a highly reflective white matte paint. A flash head (B), affixed to the outside of the dome at 90° to the viewing porthole, illuminates the interior surface with brief flashes of light. The spherical dome internally reflects light and ensures that all areas of the retina are illuminated equally. The light stimulus can be varied in both intensity and chromaticity using combinations of neutral density and chromatic filters placed in the filter slots (C) directly below the flash head or by interchanging the flash

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FIGURE 14C.2 Major components of an ERG system (see text for details).

head itself. In addition, a steady light can be projected onto the interior surface to light-adapt the retina when studying cone-mediated function (see below). A heated water bed (D) is used to maintain the mouse’s body temperature. A variety of systems for delivering light to the eye of the mouse have been described, including light-emitting diodes (LEDs), fiber-optic systems, Maxwellian view systems, and integrating spheres like that shown in Figure 14C.2. In general, the type of stimulus that is used depends on the specific questions being asked and the level of precision required. For example, fiber-optic and Maxwellian view systems provide more control over the amount of light that penetrates the eye. These techniques are useful when one is interested in knowing precisely how many photons of light are stimulating photoreceptors. However, for most applications, this level of precision is not required. For most screening purposes, a clearly detectable response to a single stimulus is sufficient to conclude that the retina is functional. In our laboratory, ERGs are recorded from the corneal surface using a gold loop electrode referenced to a similar gold wire in the mouth (Figure 14C.3). Different types of electrodes have been used including platinum, silver, copper, and even a miniature bipolar contact lens, similar to that used in humans. Most work well, but the gold and platinum wires tend to produce minimal photovoltaic artifacts at higher intensities. It is also best to use the same type of material for both the active and reference electrodes to avoid impedance mismatching. A needle electrode inserted in the tail serves as a ground. In our laboratory, signal processing is performed with custom software that allows signal averaging for the weakest signals and a signal rejection window to eliminate electrical artifacts. Isolating Rod- and Cone-Mediated Responses The mouse retina is dominated by rod photoreceptors with peak sensitivity at 510 nm corresponding to the spectral absorption characteristics of rhodopsin. Estimates of cone percentages in the mouse retina range from 1 to 10%, with newer techniques suggesting that approximately 3% of photoreceptors are

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FIGURE 14C.3 Eye electrode placement for ERG testing in the mouse.

cones.25-27 Morphologically, the cones of the mouse are indistinguishable from those of higher mammals.27 The mouse retina does not have an area centralis (i.e., an area of concentrated cones) or a visual streak. After examining the posterior pole, equator, and periphery, Carter-Dawson and LaVail27 concluded that the cone concentration was about 3% in all areas. Molecular biological, histological, and flicker electroretinographic results have established that mice have two cone photopigments, one peaking near 350 nm—ultraviolet (UV)-cone pigment—and a second near 510 nm—midwave (M)-cone pigment.28-30 ERG techniques for isolating the action spectra and absolute sensitivities of the UV-cone and M-cone-driven signals have been described.31 The properties of the cone-driven light-adapted murine ERG have also been described32,33 as have regional variations in cone function.34 Rod-mediated ERGs can be recorded using brief flashes of short-wavelength (blue) light presented to the dark-adapted eye. In our laboratory we use white light filtered through Kodak Wratten filters (W47A: λmax = 470 nm; or W47B: λmax = 449 nm) to generate the blue light required to stimulate rods. Since the spectral sensitivity of rods and the longer-wavelength cones are virtually identical, shifting the peak emission of the stimulus to shorter wavelengths minimizes the contribution from M-cones even in the dark-adapted state. Cone-mediated responses can be obtained with white flashes on a rod-saturating background. However, when the light source does not emit a significant amount of UV light, then the response to white light, presented on a rod-saturating background, is mediated primarily by the middle-wavelength sensitive cone (M-cone). Representative rod- and cone-mediated ERGs from a normal C57BL/6J mouse are shown in the left and right panels of Figure 14C.4. Each trace shows the response to a different light intensity, which is varied in 0.3 log unit steps. The b-wave of the rod-mediated response increases in amplitude, and implicit times (time from flash onset to peak of b-wave) are shortened with increasing intensity (compare heavy solid lines). The b-wave amplitude vs. intensity (I-R) series for the rod-mediated responses is summarized in the inset. Note that the b-wave amplitude saturates at the highest intensities. (The I-R series can be fitted with a Naka–Rushton function to obtain the maximum saturated b-wave amplitude, Vmax, the semisaturation intensity, k, and the ERG threshold

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Representative Rod and Cone Responses from a Normal C57BL/6J Mouse 400

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FIGURE 14C.4 Rod (left panel) and cone (right panel) ERGs obtained from a normal C57BL/6J mouse. Each trace displays the response to increasing light intensities. Left inset. Peak-to-peak amplitude vs. retinal illuminance fitted with a Naka–Rushton function to obtain the maximum saturated b-wave amplitude, Vmax, the semisaturation intensity, k, and the rod ERG threshold. Right inset. Cone b-wave amplitude vs. intensity series fitted with a linear regression to derive the cone ERG threshold intensity.

intensity.) In contrast, cone-mediated responses increase in amplitude but have relatively constant timing. The I-R series for the cone responses is shown in the inset on the right. Accuracy of the ERG As an example of the use of the ERG in mice, our group recently screened hundreds of mice to refine the location of the mouse rd3 retinal degeneration gene.35 This retinal degeneration locus was previously mapped to chromosome 1, approximately 10 + 2.5cM distal to Akp1.36 Our other goal was to exclude the mouse orthologue of USH2A.37 To begin the process of refining the position of rd3, an intercross of F2 progeny was produced from an original mating of RBF/DnJ and C57BL/6J mice. Identification of the rd3 phenotype was done with the ERG. Examples of rod-mediated responses from the rd3 mutant are shown in Figure 14C.5. The left panel shows the I-R series for a normal control littermate (filled triangles) and rd3 homozygous mice at age 4 weeks (open squares), 8 weeks (open circles), and 16 weeks (open triangles). Note that the intensity–response curves shift downward and to the right for the older mice. The shift in location of the response functions is consistent with a decrease in Vmax, the maximum saturated b-wave response, and increases in the semisaturation intensity, k, and the ERG threshold intensity. The time course of retinal degeneration in the rd3 mice compared with normal littermate controls (filled circles) is shown in the right panel. The rd3 mouse gave a saturated amplitude (Vmax) of less than 10 µv at 12 weeks of age, with nondetectable ERG responses beyond 16 weeks of age. We recorded ERGs from the first 106 F2 progeny of the intercross at approximately 2 months of age, making the discrimination of affected from unaffected animals relatively easy. The following reflects the accuracy of the ERG method that we used to phenotype a particular mouse: 32 mice that were scored by the ERG as affected were independently scored as homozygous (rd3/rd3) for all markers spanning the rd3 locus, and 74 mice that were scored as normal by the ERG were independently scored as either heterozygous (57 mice) or homozygous C57BL/6J (17 mice) for

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all markers. Based on this close correspondence between the genotype and the ERG, we were able to define the flanking markers for the rd3 gene, placing it between the markers D1Mit292/D1MIT209 and D1Mit510, a distance of 1.40 ± 0.57 cM.35 In addition, ush2a, the mouse orthologue of USH2A, was excluded from this region.37 Examples of ERG Patterns in Mice As indicated above, the ERG can be dissected to reveal the sites and mechanisms of disease action in the retina. Some examples of the different patterns of ERG responses are shown in Figure 14C.6. Shown from left to right are a rod-mediated response to dim blue flash, a mixed rod and cone response to a bright white flash (maximal response), oscillatory potentials, and a cone response to bright white light on a rod-saturating background. The first row of the figure shows representative responses from a standard C57BL/6J mouse. The second row of responses come from a reeler mouse.38 Reeler mice lack reelin, which is a large extracellular protein that is located at the stop zones for different populations of migrating neurons. These mice exhibit neuronal ectopia in the cerebral cortex, hippocampus, and cerebellum. In the retina, reelin is expressed in multiple cell types, such as rod bipolar cells, a subpopulation of amacrine cells, and retinal ganglion cells. ERGs obtained from reelin-deficient mice revealed a decreased rod-driven response, significant amplitude reductions and timing delays in the oscillatory potentials (OPs) that arise from amacrine cells, a truncated b-wave on the bright flash, and relatively normal cone-mediated function. The a-wave of the ERG was relatively preserved compared to the deficits at the level of the b-wave. Overall, these results suggested moderate abnormalities of the roddriven pathway primarily of the inner retina (bipolar, Muller, and amacrine cells) with relative preservation of the outer retina (photoreceptors). Several inbred strains have been identified recently in our laboratory that have cone dysfunction and normal rods. A colony of Peromyscus maintained at The Jackson Laboratory was previously noted to have a late-onset retinal degeneration occurring at about 18 months. However, current studies indicate that there is early-onset cone degeneration with late onset of panretinal rod

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FIGURE 14C.6 Diagnostic power of the ERG. Shown are a rod-isolated ERG, the maximal response, oscillatory potentials, and cone-isolated ERGs, obtained from a normal C57BL/6J mouse and mice of three strains with known retinal abnormalities (see text for details).

photoreceptor degeneration. An example of a Peromyscus mouse with a selective disorder of the cone system is shown in the third row of Figure 14C.6. Rod responses, shown in the left panel, are robust and indistinguishable from those of normal controls. In contrast, cone-mediated responses, shown in the far right panel, were nondetectable even at the earliest ages tested.39 Row 4 of Figure 14C.6 shows data from a new mouse model of retinoschisis.40 Ophthalmological examination of these mice revealed depigmented areas in the retina. Histology shows separation of retinal layers at the photoreceptor–Muller/bipolar cell junction. Because of the disruption of signal transmission from photoreceptors to the Muller/bipolar cells, the expectation would be that the greatest ERG deficits would be apparent at the level of the b-wave. In fact, the ERG responses show characteristic waveforms typical of a number of disease entities where the primary defect lies in the inner retina. Note the barely detectable rod b-wave, the severely truncated b-wave of the bright flash, the absent OPs, and the subnormal, but relatively preserved, cone b-wave. Photoreceptor function was unaffected as indicated by an indistinguishable, and sometimes enhanced, a-wave compared with normal controls.

FACTORS AFFECTING THE ERG There are many variables that affect the ERG and standard and consistent techniques are imperative to reduce the sometimes wide variability seen in mouse ERG recordings. Improper technique strongly affects the ERG and greatly reduces the reliability and reproducibility of data. Increased variability within ERG responses decreases the ability of the test to detect differences between strains and within strains over time, particularly when these changes are subtle. Variables that can affect the ERG include the improper use of anesthetics, variations in body temperature, insufficient dilation, inadequate light or dark adaptation, and prolonged testing, all of which can lead to a decrease in

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response amplitude. Location of the electrode on the eye can alter the amplitude of a signal by as much as to 30 to 40% and a decrease in mouse body temperature by just a few degrees is associated with a virtually nondetectable ERG. Repeated flashing, commonly used in signal averaging, reduces rod-mediated (but not cone-mediated) ERG response amplitudes by about 20% at high flash intensities unless flash presentation rate is slowed to allow sufficient recovery of rod function. Anesthetics Ketamine and xylazine mixtures are commonly used to anesthetize mice for ERG studies. Other anesthetics have been used, including Avertin, urethane, and phenobarbitol. There are two major side effects of improper use of anesthetics on the ERG. First, delivery of high doses causes an immediate reduction in ERG amplitudes. This effect is usually short-lived, and after 5 min or so, ERG responses tend to stabilize.41 Continuous infusion of small amounts of anesthetic can be one method of mitigating this effect. The second side effect has more serious consequences for the ERG. At high anesthetic doses, an acute and reversible white clouding of the lens (cataract) occurs in both rats and mice. Figure 14C.7 shows a series of photographs that demonstrate this phenomenon. Note the profound clouding of the lens in the lower frames. This clouding of the lens, if sufficiently dense, can reduce the amount of light that penetrates to the retina and can reduce ERG amplitudes. Although there are a number of factors that can accelerate this clouding, such as anoxia and altered body temperature, the main cause of this phenomenon is the dehydration of the front of the eye when blinking is stopped.42,43 Lubricating the front of the eye on a regular basis with artificial tears will prevent this phenomenon from occurring.43

FIGURE 14C.7 Development of anesthetic cataracts in the mouse. Photographs were taken at approximately 1-min intervals following a single administration of a ketamine and xylazine mixture. Note the clouding of the lens that is beginning in panel 15, approximately 10 min after anesthetic delivery.

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Mouse Body Temperature Figure 14C.8 shows the relationship between body temperature and ERG amplitudes. In this particular study, a 10 to 15% decrease in response amplitude was noted with each degree of temperature loss over the range tested. If temperature is not maintained with a heating source, temperature loss is typically 4 to 5°C over 60 min (figure inset) in a room with an ambient temperature set to 24°C. This represents b-wave reductions on the order of 40 to 50%. Commercial water beds that are warmed by circulating heated water are available as are digital probes to monitor body temperature. 1.2 40

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Pupil Size ERG response amplitudes are correlated with retinal illuminance, which is the density of light incident on the retina. The unit of measurement of retinal illuminance is the troland (td), expressed in either photopic or scotopic units.44 In general, smaller pupil diameter results in lower retinal illuminance and lower ERG response amplitudes. Because mice of different strains have different eye (and pupil) sizes, it is important to record pupil diameters so that retinal illuminance can be appropriately and correctly specified. In the mouse, pupil diameter increases rapidly with dilation medication, e.g., atropine sulfate (1%) or combinations of cyclopentolate hydrochloride (0.5%) and phenylephrine hydrochloride (2.5%). The time course of pupil dilation depends on the type of medication used, but generally halfmaximal diameter is achieved after 2 to 3 min and full pupil dilation requires 10 to 15 min. Pupil dilation is maintained for at least 2 h in the dark, but exposure to bright light can reduce this time dramatically.41

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Dark and Light Adaptation ERG amplitudes are affected by adaptation level. Typically, mice are allowed to dark- or light-adapt for a period of time to increase the sensitivity of the rod or cone systems. Illustrated in Figure 14C.9 are the rod-mediated ERGs to a blue light as a function of the time in the dark. Note that the rodmediated ERG is nondetectable with inadequate dark adaptation (bottom tracing), but grows steadily with an increase in the time in the dark. Rod-mediated responses tend to stabilize after approximately 30 to 40 min of dark adaptation. Cones also require a period of light adaptation to reach maximum output. However, unlike the rod system, the cone system reaches peak output rapidly, usually within 5 to 10 min.32,33,41

FIGURE 14C.9 Change of rod-mediated ERG response to a fixed stimulus with level of dark adaptation.

SPECIALIZED ERG RECORDING TECHNIQUES By setting specific stimulus conditions, the ERG can be used to index the functional status of a wide range of cell types and can provide information leading to a better understanding of the site and mechanisms of disease action (see Figure 14C.4). Long duration stimuli have been reported suitable for dissecting the contribution of “on” and “off” bipolar cells to the photopic ERG.45 For example, while the photopic b-wave is largely generated by cones and the depolarizing “on” bipolar cells, the activity of the hyperpolarizing off bipolar cells can limit the size and shape of the b-wave. These different components can be evaluated separately with long-duration flashes that produce distinct waveform components at flash onset and offset. Although standard ERG recordings are in response to brief flashes less than 10 ms in duration, the separation of “on” and “off” components requires longer flashes that are typically 100 to 200 ms in duration. Clinical application of the long-duration stimulus to such disease entities as congenital stationary night blindness46 and paraneoplastic night blindness47 have been reported in humans.

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a-Wave Analyses: Studies of Activation and Inactivation Steps of Phototransduction Photoreceptor structure and function can be studied by analyzing the leading edge of the ERG (called the a-wave) obtained in response to bright flashes.48-52 Prior research suggests that current quantitative rod models have the potential to discriminate structural from functional abnormalities as the underlying mechanism of disease action in retinal disease (see, for example, References 53 and 54). These techniques have been used extensively in the mouse,20,55-62 however, this type of a-wave analysis requires stimulus intensities that are substantially higher than are available with conventional photic stimulators. Intensities that clearly saturate the a-wave of the ERG are required. High output xenon arc lamps and photographic flash heads can be adapted for this purpose and can provide intensities 2.0 to 4.0 log units higher than a standard flash. An example of recordings to high-intensity flashes is shown in Figure 14C.10 (left panel) for a normal mouse. Dark-adapted ERGs were recorded to blue light flashes up to about 3.1 log scot td-s in 0.3-log unit steps. The first 30 ms of each of the responses is shown in the figure. Note that the amplitude of the a-wave is fairly stable at the highest intensities and that the time to the peak of the a-wave is shortened. The leading edge of the rod a-waves was fitted with a model of the activation phase of phototransduction.53 The fit of the model to the raw data is indicated by the dotted lines. Generated by the model are three parameters; S, RmP3, and td. S is a sensitivity parameter that scales flash energy. In general, any factor that decreases quantal catch, or affects the gain at one or more of the steps involved in phototransduction, will result in a reduction in the estimate of S. RmP3 is proportional to the magnitude of the circulating current in the rod outer-segment membrane at the time of flash presentation.48,49,51,52 A number of factors can affect this circulating current, including the ionic driving force within the cell (perhaps determined by the number of mitochondria), the electrical resistance or leakage of the photoreceptor layer, immaturity in membrane proteins that mediate the permeability of the outer limiting membrane, or the density of light-sensitive channels distributed along the rod outer segment (ROS). The parameter, td, is a brief delay before response onset.

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Prior research has reported abnormalities in S in rhodopsin mutations where the phototransduction cascade is known to be abnormal, but not in mutations where outer-segment structure is affected, as in peripherin/rds.55 An example of rod-isolated a-waves from an 8-week-old rd3 mouse are shown in Figure 14C.10 (right panel). Note that while the responses are smaller and noisier than for the normal mouse on the left, there is sufficient response to perform the a-wave analysis. For this particular mouse, RmP3 is reduced (see Figure 14C.10, right panel), consistent with shortened outer segments, but S is within normal limits, suggesting that the phototransduction cascade is unaffected. The kinetics of recovery from bright flashes can be studied using a two-flash technique. Recovery cannot be measured directly in the ERG because of the intrusion of postreceptor components. However, recovery can be inferred from the amplitude of the a-wave response to a second saturating test flash. An example of rod a-wave responses to a test flash at varying interstimulus intervals (ISIs) following a bright conditioning flash is shown in Figure 14C.11 (left panel). The test flash response in isolation is shown as the tracing labeled baseline. The other tracings show the a-wave response to the same test flash but with different ISIs ranging from 50 to 250 ms. Note that the a-wave amplitude increases as the ISI is elongated, consistent with rod functional recovery. Repetition of this two-flash paradigm with variations of the interval (ISIs) between the first and subsequent saturating flash allows determination of the recovery time course for a given conditioning flash intensity and Tc, the critical delay before the onset of recovery. Examples of normalized a-wave amplitudes to a test flash at varying ISIs are shown in Figure 14C.11 (right panel) for a dim and a bright conditioning flash (first flash). Note the faster recovery to baseline for the dim conditioning flash. We have previously reported that patients with retinitis pigmentosa and a Pro23His rhodopsin mutation not only had a decrease in the gain of activation, but also had significantly slower recovery times to bright saturating flashes.54 A modification of this technique can be used to obtain the full time course of the rod response in vivo to test flashes of subsaturating intensity.63 Although many of the specialized ERG recording techniques just described have been used extensively with humans, they have not been used widely with mice. However, as in human work, the data obtained from these recordings can provide important information about normal retinal physiology in the mouse and can provide important insights about the underlying mechanisms of disease action in mouse strains with retinal degeneration.

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FIGURE 14C.11 Kinetics of recovery to bright saturating flashes. Left panel. Representative a-wave responses to a probe flash at varying times following a bright saturating conditioning flash. Right panel. Normalized a-wave amplitudes to a test flash at varying ISIs for a dim and a bright conditioning flash (see text for details).

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The Multifocal Electroretinogram (mERG) In the mERG recording technique, small areas of the retina are stimulated simultaneously and local contributions to a massed electrical potential are extracted from a continuously recorded ERG.64-67 In contrast to the standard full-field ERG described above, which reflects the massed response of the retina to light, the mERG can provide information about the functional status of many relatively small areas of the retina. In humans, the multifocal ERG has become an increasingly popular research tool to map local cone-mediated function in a variety of retinal disorders.68-74 Our laboratory has recently extended the multifocal recording technique to map local rod-mediated function in the mouse. In a series of experiments, which are the first of its kind, we have demonstrated that the technique is feasible for use in mice,75 generating local responses (see below) that are well defined and reliable. A typical mERG stimulus is shown in Figure 14C.12. The stimulus consists of an array of hexagons displayed on a high-resolution video monitor. Each hexagon in the array is luminance-modulated according to a binary m-sequence and the local retinal response is computed as the cross-correlation between the m-sequence and a continuously recorded massed ERG. An example of a typical rod-mediated response array obtained from a normal mouse is shown in the left panel of Figure 14C.13. For this array, each stimulus element subtended approximately 5.6° × 6.9°. Note that the mERG responses are approximately uniform across the field and, although substantially smaller, have the same form as rod-mediated responses that are recorded in conventional full-field ERGs (see Figure 14C.4). The local response amplitudes decrease systematically as the size of a stimulus element is reduced (left to right) and for the smallest stimulus elements, the local responses are not well defined and difficult to discriminate from noise. For more details on this technique in mice, the reader is referred to Nusinowitz et al.75 Extending the mERG to the mouse is an important development as it allows investigation of localized changes in function of the retina. This significantly advances the quantification of retinal function by allowing investigation of regional differences in a wide variety of mouse (and more generally, rodent) models. For the testing of therapeutic interventions where localized beneficial effects can occur, as in a region of the retina receiving a gene therapy or a transplant, the technique has the potential to discover the viability of interventions that conventional ERGs would miss. Numerous technical difficulties need to be resolved before the mERG can be used routinely in mice. Because of the optics, small size of the mouse eye, and internal light scatter, there is still a question of the spatial resolution of this technique. Work is in progress in our laboratory, but even with some light scatter, preliminary data suggest that it will be possible to emphasize at least a quadrant or region of the mouse retina. The mERG technique requires clear optical media and alterations in 38 cm Radius = 15 cm

FIGURE 14C.12 Schematic representation of the multifocal ERG stimulus used for mice.

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FIGURE 14C.13 Multifocal ERGs recorded from a C57BL/6J mouse for three different local stimulus sizes (see text for details.)

body temperature, changes in level of anesthetic, and dilation medication, which, if used improperly, can all result in the development of cataracts that will reduce the local nature of a response. In addition, a technique for specifying the precise retinal locations that are stimulated, either by developing a positioning system for the mouse body, head, and eye with respect to the multifocal display or with simultaneous imaging systems that allow visualization of the retina, needs to be established.

VISUAL EVOKED CORTICAL POTENTIALS (VECP) The VECP is a gross electrical potential recorded from the visual cortex in response to a visual stimulus. That is, a visual stimulus results in the excitation of many cells in the cortex and the summed activity of these cells is recorded as the VECP on the scalp. Two anatomical constraints determine the location in the visual field from which a VECP can be recorded. First, the location of the visual cortex with respect to the surface of the skull and, second, the number of cortical cells devoted to a given region of the visual field. Because of the cortical magnification factor (M) and the anatomy of the visual cortex, the majority of the VECP response is from the central visual field in humans. The M determines the number of cells in the visual cortex devoted to analyzing a specific area of the visual field. The M is the linear extent of cortex in millimeters corresponding to 1° of visual angle. In animals with retinas that have a concentrated area of cones, the M is the greatest where the cone concentration is the highest and decreases with eccentricity. In humans, M at the fovea is 5.6 mm/° and at 10° from the fovea, the M is 1.5 mm/°.76 In mice, the cortical magnification factor is calculated to be about 6°/100 µm of cortex or 0.016 mm of cortex/° of visual field.77,78 Wagor et al.79 demonstrated that the M in the mouse is the highest near the vertical midline and decreased by a factor of 2 at 30° from the midline (i.e., from 0.027 to 0.013 mm/°). They suggested that this resulted from a higher concentration of ganglion cells devoted to the visual field near the midline. This could result in more cortical space being devoted to the central visual field in the mouse. Cortical anatomy can also play a role in the VECP response. In humans, the peripheral visual field is found at deeper locations in the calcarine fissure. The central visual field projects to the most posterior aspect of the striate cortex. This is the area closest to the scalp and, thus, electrodes placed over this area would be more effective at recording activity from the central visual field. In mice, the striate cortex (area 17, or V1) is relatively flat and exposed on the surface of the brain. This allows for VECP recordings in mice from most areas of the visual field. Thus, in mice, the VECP depends more on M than on cortical anatomy. Because the VECP depends on the number of cells in the visual cortex responding and M is the greatest near the vertical midline, the VECP is principally a function of the central visual field in the mouse.

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The Mouse Visual Pathway The retina of the mouse has from 48,000 to 65,000 ganglion cells.77,80 Higher primates have several different anatomical subclasses of ganglion cells (e.g., alpha, beta, and gamma), and it has been suggested that mice have different classes of ganglion cells based on their cell body size.77 The receptive fields of the retinal ganglion cells of the mouse have the classic center-surround arrangement found in other mammals. The size of the mouse ganglion cell receptive field is more than a log unit larger than that of the primate. This would suggest that the visual acuity of the mouse is more than a log unit less than that of primates. About half of the retinal ganglion cells project to the ipsilateral lateral geniculate nucleus (LGN) in humans. In the mouse, 2.6% of the total ganglion cell axons project to the ipsilateral lateral geniculate nucleus of the thalamus.77,81 Thus, over 97% of the axons decussate at the optic chiasm in the mouse. However, those ganglion cells with receptive fields located in binocular regions of the visual field (i.e., the central 30 to 40°) had a somewhat higher percentage (approximately 9%) of axons that remained ipsilateral. The ipsilaterally projecting axons demonstrate a divergence of connections in the LGN and occupy 14 to 18% of its volume.82 The divergence of the axons at the level of the LGN may be responsible for the amplitude of the VECP from the ipsilateral eye being as much as half that of the contralateral eye.78 As in higher mammals, the projection onto the LGN follows a retinotopic arrangement.83 Geniculocortical afferents project from the LGN to the primary visual cortex or area V1.79,84,85 Area V1, like the LGN, has a retinotopic arrangement.79,84,86,87 Thus, two cells with receptive fields near one another in the visual field will be located next to one another in the striate cortex. Single cell microelectrode studies have indicated that, as the midline is approached in area V1, the cells receptive fields move temporally in the visual field. Additionally, the superior visual field is represented posteriorly in area V1. The lateral one third of area V1 receives binocular input from the central 30 to 40° of the upper portion of each visual hemifield. Several studies have delineated the extent of area V1 in the mouse.77,79,84,86-88 Drager84 showed a map of area V1 with relation to external landmarks of the skull. Area V1 is located at the most posterior edge of the cerebral cortex. It is about 2 mm wide and 1.5 to 2.0 mm in the AP direction. The medial edge of area V1 is about 2 mm from the sagittal suture. Surrounding area V1 is area V2. All published maps of area V1 agree with Drager.79,86-88 All of the physiological classes of striate cortical cells that have been identified in cats and monkeys (i.e., center-surround cells, simple cells, complex cells, hypercomplex cells) have also been observed in the mouse.84,89,90,120 The receptive field properties of these cells are similar in the different species. Based on single cell recordings, extrastriate areas have also been identified in the mouse cortex. Wagor et al.79 identified two extrastriate areas (V2 and V3) having complete or near complete representations of the contralateral visual field. However, the cortical map that he displayed (figure 8 in Wagor et al.79) indicates that areas V2 and V3 are considerably smaller than area V1. Thus, their contribution to the VECP is limited. Based on the mouse anatomy, a normal VECP requires a normal central visual pathway from the retina to the striate cortex. VECP Stimuli Several different kinds of stimuli have been employed to produce the VECP. The stimulus chosen will depend on the question being asked. Stimuli can consist of flashes of light, checkerboard patterns, square wave gratings, or sine wave gratings.

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Flash VECP The VECP recorded in response to a flash of light can be a complicated waveform consisting of several negative- and positive-going waves.91 The flash VECP response will depend on the flash rate, intensity, and color of the stimulus. These waves have been given many different names by different investigators.92 For our purposes, the waveforms will be labeled P or N for positive- or negative-going waveforms. Additionally, each waveform will be numbered so that the first positive waveform is labeled P1, the second P2, and so on. The most consistent component of the human flash VECP is a positive waveform occurring between 100 and 150 ms after the flash of light.93 A typical flash VECP response recorded in our laboratory from a light-adapted mouse is displayed in Figure 14C.14. Each trace is the recording obtained for a different flash intensity. The mouse was anesthetized with a mixture of ketamine/xylazine (15 µg/g body weight/7 µg/g body weight) injected IP. The active electrode was a stainless steel bolt implanted 3 mm lateral to the lambda and the tip of the bolt rested on the dura. The reference electrode was a gold wire placed against the roof of the mouth and the ground was a needle under the skin near the tail. The mouse was placed in a stereotaxic (Stoelting) apparatus that held the snout. The stereotaxic apparatus was placed in a ganzfeld and the stimulus was produced by a photostimulator set at a temporal frequency of 1 Hz. Each waveform is the average of 100 repetitions. The latency (time from the flash of light to the peak of the waveform in milliseconds) of P1, N1, and P2 peaks and the amplitude of the response (the size of the waveform in microvolts from the peak of N1 to the peak of the P2) are typically determined. In this example, the latencies of the P1, N1, and P2 waves for the response to the most intense flash (top tracing) are approximately 54, 75, and 96 ms, respectively. The amplitude from N1 to P2 is 82 µv. As the flash intensity is decreased (top to bottom), the amplitude of the response decreases and the latency of peak components increases.

FIGURE 14C.14 Flash visually evoked cortical potential obtained from a normal C57BL/6J mouse (see text for details).

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Woolsey94 made early recordings of the flash VECP in mice. A craniotomy over the visual cortex was made and recordings were obtained from several locations on the surface of the dura. The stimulus was a strobe light flashing at a rate of 1 Hz. With this technique, it was possible to determine the general outline of the visual cortex (figure 4 of Woolsey94). Later studies utilized the flash VECP in mice to determine the effects of various drugs on the cortex,95-97 to examine the effects of albinism on the visual pathways,95,98 examine the effects of aging on the cortex,99 determine absolute light sensitivity,100 and to examine ultradian rhythms.101 A recent study has examined the effect of various stimulus parameters on the flash VECP response.102 In humans, many investigators have attempted to determine the neural source of each component comprising the flash VECP response. Initially, investigators determined which waveforms were generated at different levels of the visual pathway. Very early components of the flash VECP in humans (less than 65 ms after the flash onset) may be from the electroretinogram.103 Later components may be from the lateral geniculate nucleus, striate cortex, and possibly extrastriate cortex. Principal component analysis has suggested that the P1 component may be generated in area 17, the striate cortex.104 Thus, lesions of the visual pathway from the retina to the striate cortex can alter the flash VECP response (amplitude and latency). In summary, early studies of the flash VECP in mice have demonstrated the basic waveform. Recently, the flash VECP has been used to assess physiological functions of the mouse visual pathway. Future studies are needed to assess the effects of various stimulus attributes (e.g., background and stimulus intensity, stimulus wavelength, and stimulus temporal frequency) so that a complete picture of the flash VECP response can be obtained. Additionally, studies should be carried out to determine the optimum recording technique (e.g., electrode locations and anesthesia) for the mouse flash VECP. The electrode configuration and the anesthetic employed have a significant effect on the shape and timing of the VECP waveform. These studies will be necessary before the flash VECP can be used routinely to assess the mouse visual pathway. Pattern VECP The pattern VECP (i.e., the evoked response to a checkerboard pattern, sine wave, or square wave grating) can be used to assess several aspects of the visual system. The field size, pattern size, contrast, retinal location, and rate of stimulus presentation all affect the response. Thus, with the pattern VECP, visual acuity, contrast sensitivity, and motion sensitivity can be determined. There is a good correlation between these parameters determined psychophysically and electrophysiologically. The waveform for the pattern VECP has a simpler morphology than the flash VECP. A typical pattern VECP recorded from a mouse in our laboratory is displayed in Figure 14C.15. The electrode positions are the same as those for the flash VECP in Figure 14C.14. The stimulus is a checkerboard

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pattern reversing (square wave) at a temporal frequency of 4 Hz. Each check subtended an angle of 6.2° at the mouse eye. The pattern VECP consists of a negative wave at about 30 ms, a positive wave at about 60 ms, and a second negative wave at about 110 ms. The amplitude from the first negative wave to the positive wave is 6.9 µv. In humans, the positive wave occurs at about 100 ms and is often referred to as P100. Barrett et al.105 concluded that the P100 was generated in area V1 of humans. Several different pattern stimuli have been used to assess the mouse visual system.78,102,106 A number of factors have been assessed including visual acuity, cortical magnification factor (M), ocularity, contrast threshold, temporal tuning function, motion sensitivity, and luminance effect with the pattern VECP.78 The stimuli were horizontal sine wave gratings of different spatial frequencies that covered 81° × 86° of the visual field. The mice were anesthetized with 20% urethane (Sigma, 8 ml/kg) and mounted in a stereotaxic apparatus. A craniotomy was made over the visual cortex and the dura was left intact. The electrode (a resin-coated microelectrode) was placed approximately 3 mm lateral to the lambda (the intersection between the sagittal and lambdoid sutures). This corresponds to the binocular area of the striate visual cortex.84 The acuity obtained with the pattern VECP was similar to previous psychophysical measures of acuity in the mouse. The pattern VECP amplitude was plotted for a range of stimulus spatial frequencies. Peak responses were obtained with stimuli of 0.06 to 0.1 c/°. The acuity was determined by extrapolation of the high spatial frequency data to the X-axis or zero amplitude. An average acuity of 0.6 c/° was found with the pattern VECP.78 Studies of optokinetic nystagmus have suggested that the acuity of mice is about 0.5 c/°.107 Forced-choice psychophysical techniques have also resulted in acuity estimates of 0.5 to 0.6 c/°.108,109 A modification of the pattern VECP is the sweep VECP. The sweep VECP technique was developed to obtain visual acuity estimates in humans rapidly.110,111 This technique utilizes sine wave or square wave gratings. Several different pattern sizes, centered on the subject’s visual acuity, are presented in rapid succession and the individual responses are partitioned out based on the stimulus spatial frequency. A plot of spatial frequency vs. response amplitude is then obtained. Visual acuity can then be determined from this plot. Thus, this technique can be used to estimate visual acuity much more quickly than a pattern VECP technique. At present, there are no publications of sweep VECP responses in mice. Figure 14C.16 displays a sweep VECP response for a mouse in our laboratory. The electrode position and anesthesia are the same as previously described for our laboratory. Spatial frequency is plotted on the horizontal axis and the response amplitude (the second harmonic of the discrete Fourier transform) is plotted on the vertical axis. The error bars are the 95% confidence intervals. The stimulus was a horizontally oriented sine wave grating. The sweep consisted of 11 spatial frequencies (0.075, 0.09, 0.105, 0.12, 0.135, 0.15, 0.188, 0.24, 0.30, 0.47, and 0.60 cpd).

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The stimulus contrast was 80% and the temporal reversal rate (square wave) was 4 Hz. The screen luminance was 100 cd/m2 and the screen subtended 100° (H) by 82° (V) at the mouse eye. Each spatial frequency was presented for 1 s so that the entire sweep took 12 s (11 spatial frequencies + 1 second pre-adaptation). The figure displays the average of 25 sweeps. Thus, these data were obtained in 300 s or 5 min (25 sweeps × 12 s/sweep = 300 s). Stimulus production and data collection were carried out with the Enfant (Neuroscientific Corp., Farmingdale, NY) system. The data in Figure 14C.16 displays two peaks in the sweep VECP function; one peak at 0.105 cpd and a second peak at 0.30 cpd. This double-peaked function has been observed in human sweep VECP data.112,113 In humans, it has been postulated that the double peak results from the interaction of two parallel channels (i.e., the proposed transient and sustained channels) of information flow reaching the cortex at different times. Thus, at intermediate spatial frequencies (0.12 to 0.188 cpd in the mouse) they interact destructively to produce a decrease in the response amplitude. Acuity can be determined by fitting a line to the high spatial frequency data and extrapolating this line to the X-axis. Using this technique for this set of data (the dashed line in Figure 14C.16), the acuity was estimated to be 0.83 cpd. This corresponds to an acuity of 20/723. This is similar to previous reports of acuity measured with the pattern VECP and using psychophysical techniques. Contrast sensitivity has not been measured psychophysically in the mouse; however, pattern VECP data suggests that the peak contrast threshold is about 5%.78 This is similar to the peak contrast threshold of rats.114 Temporal tuning functions for the mouse striate cortex determined from pattern VECPs suggest that there is a peak at 2 to 4 Hz.78,102 On either side of this, the sensitivity is less. The temporal frequency cut off is about 12 Hz. The peak of the temporal tuning function correlates with a stimulus velocity of about 67°/s. The temporal frequency cutoff correlates with a stimulus velocity of about 200°/s. These values agree well with the optimal stimulus velocities for the cortical cells in mice.84 Drager84 found that mouse cortical cells preferred stimulus velocities from 5 to 200°/s, with some cells as high as 1000°/s. She speculated that this was the result of the anatomy of the mouse eye. The relationship between the mouse lens and the eye’s axial length would result in a minimized image of the world on the retina. Thus, only rapidly moving objects would optimally stimulate retinal cells. By driving an electrode to various layers of the cortex and monitoring the pattern VECP, the source of the VECP can be determined. On either side of the source, the VECPs will have opposite polarities. It has been determined that the source of the pattern VECP in the mouse is in the supragranular layers of area V1.78 This agrees with work on the rat 115,116 and monkey.117 In conclusion, the pattern VECP has been used to assess several aspects of the mouse visual system. The pattern VECP results correspond to the psychophysical findings for the mouse and can be used to make these measurements more efficiently. However, since there are only two publications at the time of this writing that have demonstrated pattern VECPs in the mouse, these results need to be replicated and the optimal recording conditions need to be identified. Multifocal VECP (mVECP) The mVECP has been recorded in humans.118,119 The technique is virtually identical to that described for the mERG (see above) except that local VECP responses are determined from a continuously recorded EEG. There are no publications of multifocal VECP recordings in mice. Preliminary data from our laboratory suggest that the technique is feasible in mice and may prove to be a useful technique for localization of visual field defects. However, our early data suggest that the local stimuli must be relatively large to obtain reliable and repeatable waveforms, so that the spatial resolution is poorer than that which we could achieve with the mERG (see above).

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FIGURE 14C.17 Local VECP responses obtained from a representative normal C57BL/6J mouse (see text for details).

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Figure 14C.17 displays the results of multifocal VECP recordings from a C57BL/6J mouse in our laboratory. The stimulus, a seven hexagon display, was placed at 30 cm from the eye. Each local target subtended approximately 18° × 17°, substantially larger than the 5.6° × 6.9° elements used for the mERG. Each hexagon in the array was luminance-modulated according to the binary m-sequence. The same m-sequence was used for all hexagons in the display. The stimulation rate was controlled by placing blank frames (N = 19) between successive flashes in the m-sequence. This resulted in the minimum interflash time interval being 252.7 ms (13.3 × 19). The m-sequence length was 210 –1, which required 4 min 19 s to complete. Data collection and analysis was carried out with the VERIS system. The VECP response to each hexagon in the display is shown in Figure 14C.17. Note that while the individual responses are more variable, they have the same basic shape as those obtained with the full-field flash VECP (see Figure 14C.14). The individual hexagon responses have a negative peak at about 64 ms and a positive peak at about 83 ms. The identification of peak components and the correspondence with the full-field flash are more evident in Figure 14C.18, which is the sum of the individual tracings shown in Figure 14C.17. Thus, multifocal VECPs can be recorded in mice. This may allow for an evaluation of discrete areas of the visual field in mice. However, before these can be used routinely in mice, various stimulus parameters must be investigated to optimize the response.

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GENERAL CONCLUSIONS The electrophysiological techniques described in this section can be powerful tools to better understand the sites and mechanisms of disease action in mouse models of ocular disease. The ERG is a commonly used technique to assess panretinal function and can be dissected to quantify and evaluate the functional integrity of the outer and inner retinal layers. In contrast to the conventional ERG, which represents the summed activity across the retina, the mERG has been demonstrated to be a useful technique for isolating retinal function in small areas of the retina. Although the mERG is in development with respect to mouse applications, it should be a useful addition to the battery of tests to describe retinal phenotype and to evaluate the efficacy of certain therapeutic interventions. At the present time there are no internationally accepted standards for recording ERGs in mice. It is strongly recommended that standards be adopted so that comparisons of responses between laboratories can easily be made. It is clear that VECPs are readily recorded from the central visual field of mice. The flash VECP can be employed to determine if there are any lesions of the visual pathway from the retina to the striate cortex. Pattern VECPs can be used to assess several aspects of the mouse visual system, including visual acuity, cortical magnification factor (M), ocularity, contrast threshold, temporal tuning function, motion sensitivity, and luminance effects. Multifocal VECPs may prove beneficial in assessing specific aspects of the visual field in mice. However, future studies are needed to extend this work so that the effect of various stimulus parameters on the VECP in mice is better understood. As with the ERG, there are no current standards for recording VECPs in mice. A standard protocol for mice VECPs is recommended.

REFERENCES 1. Granit, R., Two types of retinae and their electrical responses to intermittent stimuli in light and dark adaptation, J. Physiol., 85:421, 1935. 2. Granit, R., Receptors and Sensory Perception, Yale University Press, New Haven, CT, 1955. 3. Penn, R.D. and Hagins, W.A., Signal transmission along retinal rods and the origin of the electroretinographic a-wave, Nature, 223:201, 1969. 4. Heynen, H. and Van Norren, D., Origin of the electroretinogram in the intact macaque eye. I., Vision Res., 25, 697, 1985. 5. Miller, R.F. and Dowling, J.E., Intracellular responses of Muller (glial) cells of the mudpuppy retina: their relation to the b-wave of the electroretinogram, J. Neurophysiol., 33:323, 1970. 6. Newman, E.A. and Odette, L.L., Model of electroretinogram b-wave generation: a test of the K+ hypothesis, J. Neurophysiol., 51:164, 1984. 7. Robson, J.G. and Frishman, L.J., Response linearity and kinetics of the cat retina: the bipolar cell component of the dark-adapted electroretinogram, Vis. Neurosci., 12:837, 1995. 8. Stockton, R.A. and Slaughter, M.M., B-wave of the electroretinogram: reflection of ON bipolar cell activity, J. Gen. Physiol., 93:101, 1989. 9. Xu, X.J. and Karwoski, C.J., Current source density analysis of retinal field potentials. 2: pharmacological analysis of the b-wave and m-wave, J. Neurophysiol., 72:96, 1994. 10. Heynen, H., Wachtmeister, L., and Van Norren, D., Origin of the oscillatory potentials in the primate retina, Vision Res., 25:1365, 1985. 11. Karwoski, C. and Kawasaki, K., Oscillatory potentials, in Handbook of Clinical Electrophysiology of Vision Testing, Heckenlively, J.H. and Arden, G.B., Eds., Mosby Year Book, St. Louis, 125–128, 1991. 12. Ogden, T.E., The oscillatory waves of the primate electroretinogram, Vision Res., 13:1059, 1973. 13. Wachtmeister, L., Oscillatory potentials in the retina: what do they reveal, Prog. Retinal Eye Res., 17:485, 1998. 14. Wachtmeister, L. and Dowling, J.E., The oscillatory potentials of the mudpuppy retina, Invest. Ophthalmol. Vis. Sci., 17:1176, 1978.

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15. Griff, E.R., Electroretinographic components arising in the distal retina, in Principles and Practice of Clinical Electrophysiology of Vision, Heckenlively, J.R. and Arden, G.B., Eds., Mosby, St. Louis, 1991, 91–98. 16. Sieving, P.A., Frishman, L.J., and Steinberg, R.H., Scotopic threshold response of proximal retina in cats, J. Neurophysiol., 56:1049, 1986. 17. Bush, R.A., Hawks, K.W., and Sieving, P.A., Preservation of inner retinal responses in the aged Royal College of Surgeons rat. Evidence against glutamate excitotoxicity in photoreceptor degeneration, Invest. Ophthalmol. Vis. Sci., 36:2054, 1995. 18. Sugawara, T., Sieving, P.A., and Bush, R.A., Quantitative relationship of the scotopic and photopic ERG to photoreceptor cell loss in light damaged rats, Exp. Eye Res., 70:693, 2000. 19. Green, D.G. et al., Electrophysiology and density of retinal neurons in mice with a mutation that includes the Pax2 locus, Invest. Ophthalmol. Vis. Sci., 38:919, 1997. 20. Peachey, N.S. et al., Functional consequences of oncogene-induced horizontal cell degeneration in the retinas of transgenic mice, Vis. Neurosci., 14:627, 1997. 21. Marti, A. et al., Light-induced cell death of retinal photreceptors in the absence of p53, Inv. Ophthal. Vis. Sci., 39:846, 1998. 22. Ruether, K. et al., Retinoschisis-like alterations in the mouse eye caused by gene targeting of the Norrie disease gene, Invest. Ophthalmol. Vis. Sci., 38:710, 1997. 23. Shaaban, S.A. et al., Transgenic mice expressing a functional human photopigment, Invest. Ophthalmol. Vis. Sci., 39:1036, 1998. 24. Smith, S.B. and Hamasaki, D.I., Electroretinographic study of the C57BL/6-mivit/mivit mouse model of retinal degeneration, Invest. Ophthalmol. Vis. Sci., 35:3119, 1994. 25. Menner, E., Untersuchungen über die Retina mit besonderer Berucksichtigung der ausseren Kornerschichte, Z. Vergl. Physiol., 8:761, 1928. 26. Sidman, R.L., Histochemical studies on photoreceptor cells, Ann. N.Y. Acad. Sci., 74:182, 1958. 27. Carter-Dawson, L. and LaVail, M.M., Rods and cones in the mouse retina. I. Structural analysis using light and electron microscopy, J. Comp. Neurol., 188:245, 1979. 28. Jacobs, G.H., Neitz, J., and Deegan III, J.F., Retinal receptors in rodents maximally sensitive to ultraviolet light, Nature, 353, 655, 1991. 29. Jacobs, G.H., Ultraviolet vision in vertebrates, Am. Zool., 32:544, 1992. 30. Sun, H., Macke, J.P., and Nathans, J., Mechanisms of spectral tuning in the mouse green cone pigment, Proc. Natl. Acad. Sci. U.S.A., 94, 8860, 1997. 31. Lyubarsky, A.L. et al., UV- and midwave-sensitive cone-driven retinal responses of the mouse: a possible phenotype for coexpression of cone photopigments, J. Neurosci., 19:442, 1999. 32. Peachey, N.S. et al., Properties of the mouse cone-mediated electroretinogram during light adaptation, Neurosci. Lett., 162:9, 1993. 33. Ekesten, B., Gouras, P., and Moschos, M., Cone properties of the light-adapted murine ERG, Doc. Ophthalmol., 97:23, 1998. 34. Calderone, J.B. and Jacobs, G.H., Regional variations in the relative sensitivity to UV light in the mouse retina, Vis. Neurosci., 12:463, 1995. 35. Danciger, J.S. et al., Genetic and physical maps of the rd3 locus; exclusion of the mouse ortholog of USH2A, Mamm. Genome, 10:657, 1999. 36. Chang, B. et al., New mouse primary retinal degeneration (rd-3), Genomics, 16:45, 1993. 37. Sumegi, J. et al., The construction of a yeast artificial chromosome (YAC) contig in the vicinity of the Usher syndrome type IIa (USH2A) gene in 1q41, Genomics, 35:79, 1996. 38. Rice, D.S. et al., The Reelin signaling pathway modulates the formation of synaptic circuitry in the retina, Neuron, 31:1, 2001. 39. Nusinowitz, S., Performing electroretinography and analysis in the mouse, presented at Retinal Degenerations Symposium, The Jackson Laboratory, Bar Harbor, ME, 1999. 40. Nusinowitz, S. et al., A mouse model of retinoschisis associated with a collagen II mutation, Invest. Ophthalmol. Vis. Sci., 40 (Suppl.):S364, 2001. 41. Nusinowitz, S., Azimi, A., and Heckenlively, J.H., Adaptation, body temperature, pupil size, and anesthetic effects on the electroretinogram (ERG) in mice, Invest. Ophthalmol. Vis. Sci., 41 (Suppl.):S495, 2000.

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42. Hockwin, O. and Koch, H.R., Arzneimittelnebenwirkungen am Auge, Gustav Fischer, Stuttgart, 1977, 49 [in German]. 43. Ridder, W.H., Heckenlively, J.H., and Nusinowitz, S., Cataractogenesis in anesthetized mice, Exp. Eye Res., submitted. 44. Wyszecki, G. and Stiles, W.S., Color Science: Concepts and Methods, Quantitative Data and Formulas, Wiley, New York, 1982. 45. Sieving, P., Photopic “ON” and “OFF” pathway abnormalities in retinal dystrophies, Trans. Am. Ophthalmol. Soc., 91:701, 1993. 46. Miyake, Y. et al., On- and off-responses in photopic electroretinogram in complete and incomplete types of congenital stationary night blindness, Jpn. J. Ophthalmol., 31:81, 1987. 47. Alexander, K. et al., “On” response defect in paraneoplastic night blindness with cutaneous malignant melanoma, Invest. Ophthalmol. Vis. Sci., 33:477, 1992. 48. Hood, D.C. and Birch, D.G., The a-wave of the human electroretinogram and rod receptor function, Invest. Ophthalmol. Vis. Sci., 31:2070, 1990. 49. Hood, D.C. and Birch, D.G., A quantitative measure of the electrical activity of human rod photoreceptors using electroretinography, Vis. Neurosci., 5:379, 1990. 50. Breton, M.E. and Montzka, D., Empiric limits of rod photocurrent component underlying a-wave response in the electroretinogram, Doc. Ophthalmol., 79:337, 1992. 51. Breton, M.E. et al., Analysis of ERG a-wave amplification and kinetics in terms of the G-protein cascade of phototransduction, Invest. Ophthalmol. Vis. Sci., 35:295, 1994. 52. Cideciyan, A.V. and Jacobson, S.G., Negative electroretinograms in retinitis pigmentosa, Invest. Ophthalmol. Vis. Sci., 34:3253, 1993. 53. Hood, D.C. and Birch, D.G., Rod phototransduction in retinitis pigmentosa: estimation and interpretation of parameters derived from the rod a-wave, Invest. Ophthalmol. Vis. Sci., 35:2948, 1994. 54. Birch, D.G. et al., Abnormal activation and inactivation mechanisms of rod transduction of patients with autosomal dominant retinitis and the PRO-23-HIS mutation, Invest. Ophthalmol. Vis. Sci., 36:1603, 1996. 55. Kedzierski, W. et al., Generation and analysis of transgenic mice expressing P216L-substituted Rds/Peripherin in rod photoreceptors, Invest. Ophthalmol. Vis. Sci., 38:498, 1997. 56. Goto, Y. et al., Rod phototransduction in transgenic mice expressing a mutant opsin gene, J. Opt. Soc. Am. A, 13:577, 1996. 57. Goto, Y. et al., Functional abnormalities in transgenic mice expressing a mutant rhodopsin gene, Invest. Ophthalmol. Vis. Sci., 36:62, 1995. 58. Goto, Y. et al., Rod phototransduction in transgenic mice expressing a mutant opsin gene, J. Opt. Soc. Am. A, 13:577, 1996. 59. Yang, R.B. et al., Disruption of a retinal guanylyl cyclase gene leads to cone-specific dystrophy and paradoxical rod behavior, J. Neurosci., 19:5889, 1999. 60. Ren, J.C., LaVail, M.M., and Peachey, N.S., Retinal degeneration in the nervous mutant mouse. III. Electrophysiological studies of the visual pathway, Exp. Eye Res., 70:467, 2000. 61. Pardue, M.T. et al., A naturally occurring mouse model of X-linked congenital stationary night blindness, Invest. Ophthal. and Vis. Sci., 39:2443, 1998. 62. Rohrer, B. et al., Role of neurotrophin receptor TrkB in the maturation of rod photoreceptors and establishment of synaptic transmission to the inner retina, J. Neurosci., 19:8919, 1999. 63. Hetling, J.R. and Pepperberg, D.R., Sensitivity and kinetics of mouse rod flash responses determined in vivo from paired-flash electroretinograms, J. Physiol., 516:593, 1999. 64. Sutter, E.E., The fast m-transform: a fast computation of cross-correlations with binary m-sequences, Soc. Ind. Appl. Math., 20:686, 1991. 65. Sutter, E.E. and Tran, D., The field topography of ERG components in man. I. The photopic luminance responses, Vision Res., 32:433, 1992. 66. Bearse, M.A. and Sutter, E.E., Imaging localized retinal dysfunction with the multifocal electroretinogram, J. Opt. Soc. Am. A, 13:634, 1996. 67. Hood, D.C. et al., A comparison of the components of the multi-focal and full-field ERGs, Vis. Neurosci., 14:533, 1997.

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68. Hood, D.C. et al., Assessment of local retinal function in patients with retinitis pigmentosa using the multi-focal ERG technique, Vision Res., 38:163, 1997. 69. Kondo, M. et al., Clinical evaluation of multifocal electroretinogram, Invest. Ophthalmol. Vis. Sci., 36:2146, 1995. 70. Kretschmann, U. et al., Electroretinographic campimetry in a patient with crystalline retinopathy, Ger. J. Ophthalmol., 5:399, 1997. 71. Kretschmann, U. et al., ERG campimetry using a multi-input stimulation technique or mapping of retinal function in the central visual field, Ophthalmic Res., 28:303, 1996. 72. Palmowski, A.M. et al., Mapping of retinal function in diabetic retinopathy using the multifocal electroretinogram, Invest. Ophthalmol. Vis. Sci., 38:2586, 1997. 73. Parks, S. et al., Functional imaging of the retina using the multi-focal electroretinograph: a control study, Br. J. Ophthalmol., 80:831, 1996. 74. Seeliger, M. et al., Multifocal electroretinography in retinitis pigmentosa, Am. J. Ophthalmol., 125, 214, 1998. 75. Nusinowitz, S., Ridder, W.H., III, and Heckenlively, J.R., Multi-focal rod electroretinograms in mice, Invest. Ophthalmol. Vis. Sci., 4:2848, 1999. 76. Rovamo, J. and Virsu, V., An estimation and application of the human cortical magnification factor, Exp. Brain Res., 37:495, 1979. 77. Drager, U.C. and Olsen, J.F., Origins of crossed and uncrossed retinal projections in pigmented and albino mice, J. Comp. Neurol., 191:383, 1980. 78. Porciatti, V., Pizzorusso, T., and Maffei, L., The visual physiology of the wild type mouse determined with pattern VEPs, Vision Res., 39:3071, 1999. 79. Wagor, E., Mangini, N.J., and Pearlman, A.L., Retinotopic organization of striate and extrastriate visual cortex in the mouse, J. Comp. Neurol., 193:187, 1980. 80. Gyllensten, L., Malmfors, T., and Norrlin-Grettve, M.L., Developmental and functional alterations in the fiber composition of the optic nerve in visually deprived mice, J. Comp. Neurol., 128:413, 1966. 81. Balkema, G.W. and Drager, U.C., Origins of uncrossed retinofugal projections in normal and hypopigmented mice, Vis. Neurosci., 4:595, 1990. 82. LaVail, J.H., Nixon, R.A., and Sidman, R.L., Genetic control of retinal ganglion cell projections, J. Comp. Neurol., 182:399, 1978. 83. Godement, P., Salaun, J., and Imbert, M., Prenatal and postnatal development of retinogeniculate and retinocollicular projections in the mouse, J. Comp. Neurol., 230:552, 1984. 84. Drager, U.C., Receptive fields of single cells and topography in mouse visual cortex, J. Comp. Neurol., 160:269, 1975. 85. Drager, U.C., Observations on monocular deprivations in mice, J. Neurophysiol., 41, 28, 1978. 86. Simmons, P.A. and Pearlman, A.L., Retinotopic organization of the striate cortex (area 17) in the reeler mutant mouse, Dev. Brain Res., 256:124, 1982. 87. Gordon, J.A. and Stryker, M.P., Experience-dependent plasticity of binocular responses in the primary visual cortex of the mouse, J. Neurosci., 16:3274, 1996. 88. Valverde, F. and Esteban, M.E., Peristriate cortex of mouse: Location and the effects of enucleation on the number of dendritic spines, Brain Res., 9:145, 1968. 89. Hubel, D.H. and Wiesel, T.N., Receptive fields, binocular interaction and functional architecture in the cat’s visual cortex, J. Physiol., 160:106, 1962. 90. Hubel, D.H. and Wiesel, T.N., Receptive fields and functional architecture of monkey striate cortex, J. Physiol., 195:215, 1968. 91. Ciganek, M.L., The EEG response (evoked potential) to light stimulus in man, Electroencephalogr. Clin. Neurophysiol., 13:165, 1961. 92. Harding, G.F., History of visual evoked cortical testing, in Principles and Practice of Clinical Electrophysiology of Vision, Heckenlively, J.R. and Arden, G.B., Eds., Mosby, St. Louis, 1991, 17–22. 93. Gastaut, H. and Regis, H., Visual evoked potentials recorded transcranially in man, in Proctor, L.D. and Adey, W.R., Eds., The Analysis of Central Nervous System and Cardio-Vascular System Data Using Computer Methods, NASA, Washington, D.C., 1965, 7–34. 94. Woolsey, T.A., Somatosensory, auditory and visual cortical areas of the mouse, Johns Hopkins Med. J., 121:91, 1967.

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95. Henry, K.R. and Rhoades, R.W., Relation of albinism and drugs to the visual evoked potential of the mouse, J. Comp. Physiol. Psychol., 92:271, 1978. 96. Adachi-Usami, E., Ikeda, H., and Satoh, H., Haloperidol delays visually evoked cortical potentials but not electroretinograms in mice, J. Ocular Pharmacol., 6:203, 1990. 97. Tebano, M.T. et al., Effects of cholinergic drugs on neocortical EEG and flash-visual evoked potentials in the mouse, Neuropsychobiology, 40:47, 1999. 98. Henry, K.R., Rhoades, R.W., and Haythorn, N.M., Effects of ambient lighting and the albino gene on the developing visual evoked potential of the mouse, Physiol. Psychol., 5:204, 1977. 99. Adachi-Usami, E., Senescence of visual function as studied by visually evoked cortical potentials, Jpn. J. Ophthalmol., 34:81, 1990. 100. Green, D.G., Herreros De Tejada, P., and Glover, M.J., Electrophysiological estimates of visual sensitivity in albino and pigmented mice, Vis. Neurosci., 11: 919, 1994. 101. Mazzucchelli, A. et al., Ultradian rhythms in the N1-P2 amplitude of the visual evoked potential in two inbred strains of mice: DBA/2J and C57BL/6, Behav. Brain Res., 67:81, 1995. 102. Strain, G.M. and Tedford, B.L., Flash and pattern reversal visual evoked potentials in C57BL/6J and B6CBAF1/J mice, Brain Res. Bull., 32:57, 1993. 103. Allison, T. et al., The scalp topography of human visual evoked potentials, Electroencephalogr. Clin. Neurophysiol., 42:185, 1977. 104. Maier, J. et al., Principal component analysis for source localization of VEPs in man, Vision Res., 27:165, 1987. 105. Barrett, G. et al., A paradox in the lateralisation of the visual evoked response, Nature, 261:253, 1976. 106. Porciatti, V., Pizzorusso, T., and Maffei, L., Vision in mice with neuronal redundancy due to inhibition of developmental cell death, Vis. Neurosci., 16:721, 1999. 107. Sinex, D.G., Burdette, L.J., and Pearlman, A.L., A psychophysical investigation of spatial vision in the normal and reeler mutant mouse, Vision Res., 19:853, 1979. 108. Gianfranceschi, L., Fiorentini, A., and Maffei, L., Behavioural visual acuity of wild-type and BC-l2 transgenic mouse, Vision Res., 39:569, 1999. 109. Prusky, G.T., West, P.W., and Douglas, R.M., Behavioral assessment of visual acuity in mice and rats, Vision Res., 40:2201, 2000. 110. Tyler, C.W. et al., Rapid assessment of visual function: an electronic sweep technique for the pattern visual evoked potential, Invest. Ophthalmol. Vis. Sci., 18:703, 1979. 111. Norcia, A.M. and Tyler, C.W., Spatial frequency sweep VEP: visual acuity during the first year of life, Vision Res., 25:1399, 1985. 112. Strasburger, H., Scheidler, W., and Rentschler, I., Amplitude and phase characteristics of the steady state visually evoked potential, Appl. Optics, 27:1069, 1988. 113. Gottlob, I. et al., Predicting optotype visual acuity by swept spatial visual-evoked potentials, Clin. Vision Sci., 8:417, 1993. 114. Birch, D. and Jacobs, G.H., Spatial contrast sensitivity in albino and pigmented rats, Vision Res., 19:933, 1979. 115. Fontanesi, G. et al., Somatostatin depletion modifies the functional activity of the visual cortex in the rat, Vis. Neurosci., 3:327, 1996. 116. Pizzorusso, T. et al., Temporal aspects of contrast visual evoked potentials in the pigmented rat: effect of dark rearing, Vision Res., 37:389, 1996. 117. Schroeder, C.E. et al., Striate cortical contribution to the surface-recorded pattern-reversal VEP in the alert monkey, Vision Res., 31:1143, 1991. 118. Klistorner, A.I. et al., Multifocal topographic visual evoked potential: improving objective detection of local visual field defects, Invest. Ophthalmol. Vis. Sci., 39:937, 1998. 119. Hood, D.C. et al., An interocular comparison of the multifocal VEP: a possible technique for detecting local damage to the optic nerve, Invest. Ophthalmol. Vis. Sci., 41:1580, 2000. 120. Mangini, N.J. and Pearlman, A.L., Laminar distribution of receptive field properties in the primary visual cortex of the mouse, J. Comp. Neurol., 193:203, 1980.

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CONTENTS Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .345 Information Resources . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .346 Genetic Resources . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .348 Future Directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .349 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .352

INTRODUCTION With the ability to manipulate the mouse genome, the use of mice in biomedical research has increased significantly over the past 20 years. Genetically engineered mice have expanded the already extensive number of inbred strains and mice carrying spontaneous or more classically induced mutations (e.g., by radiation) used to study the eye. Genetically defined mice (inbred strains and mutations carried on an inbred strain) offer a number of advantages as models to understand basic biological processes and the pathogenesis of human disease. Not only do mice share most of the same genes as humans, the genes function essentially the same way within a biological context. Mice are also manageable research tools: 1. 2. 3. 4. 5.

Because of their size and nature, they are easy to maintain in large numbers. They are easy to manipulate genetically. They reproduce quickly and often. They have a relatively short life span. They provide reproducible experimental systems for understanding normal development and function.

Finally, the use of genetically defined mice permit studies that may be inappropriate or impossible in humans. Because of their versatility and the exponential growth of the number of mouse models available to the research community, more and more scientists are adopting mice as their primary model organism. There are several challenges for the novice mouse user. Appropriate model selection involves more than choosing a strain carrying a specific mutation. The phenotypic characteristics and pathophysiology of mice carrying spontaneous or genetically engineered mutations result from a combination of the modified gene, the genetic background, and the surrounding environment. It is therefore necessary to obtain as much information as possible about a given mouse model prior to its selection and use. Numerous information resources are now available to the researcher on the Web. Repositories like the Mouse Mutant Resource (MMR) and Induced Mutant Resource (IMR) at The Jackson Laboratory and the Mouse Mutant Regional Resource Center (MMRRC) have been established to ensure the genetic quality of valuable models, to archive these resources, to standardize environmental factors, and to distribute these mice to the research community.

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INFORMATION RESOURCES The World Wide Web offers a virtual library of information resources to the scientist interested in finding out what is available to the research community. It is useful to take advantage of the as-yet limited number of relevant information warehouses and databases available (Tables 15.1 and 15.2) when trying to select a new mouse model or learn more specific information about the nature of the model (e.g., genetic and phenotypic characteristics, homology data, animal husbandry and maintenance, etc.). As part of its mission to the research community, The Jackson Laboratory is an essential knowledge center for information about mice (Table 15.2). The Mouse Genome Informatics (MGI) project was initiated in 1992 to integrate and centralize the numerous mouse-related databases to facilitate new data integration and analysis. Released on the World Wide Web in June 1994, the first version incorporated all the data sets from the Genomic Database of the Mouse (GBASE) the Mouse Locus Catalog (MLC), Mouse Linkage Database and Programs (MLDP), Homology Database and Programs (HMDP), and the Probes and PCR databases describing molecular reagents and polymorphisms (MusProb and MusPCR). Since its inception, the Web site has undergone major construction to incorporate new databases and data sets, to improve functionality and query mechanisms, and to develop and enhance analysis tools. TABLE 15.1 General Resources on Mouse Information Dysmorphic Human-Mouse Homology Database (DHMHD): http://www.hgmp.mrc.ac.uk/DHMHD/dysmorph.html

DHMHD is a composite of three separate databases of human and mouse malformation syndromes together with a database of mouse/human syntenic regions. The database can be searched by (1) specifying specific malformations, clinical features, or chromosome locations; (2) homology; (3) asking for human syndromes located at a chromosome region homologous with a specific mouse chromosome region (and vice versa, from human to mouse).

Mouse Atlas and Gene Expression Database Project: The Edinburgh Mouse Atlas, Standard Anatomical Nomenclature: http://genex.hgu.mrc.ac.uk

The U.K. MRC Human Genetics Unit in Edinburgh is developing a digital atlas of mouse development and a database to be a resource for spatially mapped data such as in situ gene expression and cell lineage. The project is in collaboration with the Department of Anatomy, University of Edinburgh. The gene expression database is being developed as part of the Mouse Gene Expression Information Resource (MGEIR) in collaboration with The Jackson Laboratory. Established in 1995, MKMD is the BioMedNet fully searchable database of phenotypic information related to knockout and classical mutations in mice.

Mouse Knockout and Mutation Database (MKMD): http://www.biomednet.com/db/mkmd Online Mendelian Inheritance in Man (OMIM): http://www3.ncbi.nlm.nih.gov/Omim

OMIM catalogs human phenotypes and genotypes and relevant mouse models. Authored and edited by Dr. Victor A. McKusick and his colleagues and developed for the World Wide Web by NCBI (the National Center for Biotechnology Information), the database contains textual information, pictures, and reference information. It also contains copious links to the NCBI Entrez database of MEDLINE articles and sequence information.

Whole Mouse Catalog: http://www.rodentia.com/wmc

This Web site serves as an information warehouse for scientific researchers using mice or rats in their work.

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TABLE 15.2 Information Resources at The Jackson Laboratory http://www.jax.org Resource

Description

JAX® Mice Database http://www.jax.org/jaxmice/pricelist

The JAX Mice Database offers a wealth of information including phenotypes, gene descriptions, prices, stock numbers, availability, genotypes, gene names, controls, and other helpful information regarding JAX mice. Search by research application (e.g., Sensorineural Research: cataracts, retinal degeneration, etc.).

Mouse Genome Informatics Project http://www.informatics.jax.org

The Mouse Genome Informatics Database provides integrated access to data on the genetics, genomics and biology of the laboratory mouse. The projects contributing to this resource are: Mouse Genome Database (MGD) Project includes data on gene characterization and nomenclature, mapping, gene homologies among mammals, sequence links, phenotypes, allelic variants and mutants, and strain data. Gene Expression Database (GXD) Project integrates different types of gene expression information from the laboratory mouse and provides an electronic index of published experiments on endogenous gene expression during mouse development. Mouse Genome Sequence (MGS) Project is integrating emerging mouse genomic sequence data with the genetic and biological data available in MGD and GXD. Mouse Tumor Biology (MTB) Database Project catalogs information relevant to using mouse models in cancer research including tumor names and classifications, tumor incidence and latency data, tumor pathology reports and images, genetic factors, and references.

Inbred Strain Characteristics Dr. Michael Festing’s Inbred Strain Characteristics is a searchable, hypertext (by Dr. Michael F.W. Festing) access listing of inbred strains of mice and rats with associated strain characteristics. through http://www.informatics.jax.or It is a valuable resource for basic characteristics of inbred strains including incidence of spontaneous diseases and which commonly used inbred strain carry mutations affecting eye development and vision. Mouse Phenome Database http://www.jax.org/phenome

An information repository for protocols and raw data related to the phenotypic characteristics of commonly used and genetically diverse inbred mouse strains. Provides current project information including opportunities for community participation in this coordinated international effort.

Transgenic/Targeted Mutation Database (TBASE) http://www.jax.org/tbase

TBASE is an information database of mice carrying transgenes and targeted mutations (“knockouts”) generated by the worldwide research community.

At present, the MGI Web site (http://www.informatics.jax.org) provides a unique integrated access to various sources for information on the genetics and biology of the laboratory mouse. It comprises the Mouse Genome Database1 (MGD), the Gene Expression Database2 (GXD), Mouse Genome Sequence (MGS) project, and related resources including the Mouse Tumor Biology (MTB) database,3 the Rat Data resource, Michael Festing’s Listing of Inbred Strains of Mice and Rats, and MouseBLAST. MGI also provides links to a wide variety of other scientific Web sites. The MGI group, as a member of the Gene Ontology Consortium (www.geneontology.org), recently implemented a new classification system for genes and gene products.4 Defined, structured vocabularies, called gene ontologies, provide an annotation pipeline across eurkaryotic species to describe the “molecular function.” “biological process,” and “cellular component,” aspects of a given gene. Current annotation data sets being developed include the following model organisms: Drosophila melanogaster, Caenorhabditis elegans, Mus musculus, Arabidopsis thaliana, and Saccharomyces cerevisiae.

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In addition to the gene ontology (GO) terms, MGI is constructing vocabularies to annotate phenotypic information on the thousands of mutant strains of mice currently listed in the Mouse Genome Database. More information about the Gene Ontology project and the use of the GO terms in MGI can be found on their Web site http://www.informatics.jax.org/mgihome/GO/ontology.shtml. The Jackson Laboratory is also the home of the recently established Mouse Phenome Database (MPD) project. The goal of MPD is “to establish a collection of baseline phenotypic data on commonly used and genetically diverse inbred mouse strains through a coordinated international effort.”5 There is a tremendous need for comprehensive phenotypic information on inbred mouse strains. The laboratory mouse is the primary genetic model for exploring normal human biology and disease. In addition, hundreds (perhaps thousands) of spontaneous and induced mutations are maintained on an inbred background; the resulting phenotype is most often combination of both the mutation and the genetic background. The MPD project will collaborate with scientists to gather phenotypic information on a diverse panel of inbred strains. The MPD Web site (http://www.jax.org/phenome) will help researchers select appropriate strains in a wide variety of research applications including physiological testing, gene identification, drug discovery, toxicology studies, mutagenesis, disease onset and susceptibility, and QTL analyses. Electronic listservers, or electronic bulletin boards, provide a valuable and often overlooked service to the research community (Table 15.3). Most listservers are geared toward a specific research area. With thousands of subscribers, a researcher can often find a source for a rare mouse model or even gather information on whether an observed phenotype is the result of the mutation or of the environment.

GENETIC RESOURCES Since its inception in 1929, The Jackson Laboratory has served as a central repository, developing and distributing a wide variety of inbred strains and mice carrying mutations. Originally held in individual research laboratories it became clear by the late 1940s that it was important to consolidate the growing number of mutant mouse strains into central colonies. Early repositories like the Mouse Mutant Stocks Center (MMSC) and the Mouse Mutant Gene Resource (MMGR) were combined to

TABLE 15.3 Electronic Listservers COMPMED: http://www.aalas.org/association/links/compmed.htm

COMPMED is a comparative Medicine Discussion List–Mailing List for discussing the topics of comparative and laboratory animal medicine.

Mouse Genome Informatics e-mail lists: MGI maintains several e-mail lists for the genetics research http://www.informatics.jax.org/mgihome/lists/lists.shtml community. The list service uses the Lyris list service software package, version 3.0. MGI-LIST: Forum for topics in mouse genetics, MGI news updates RAT-LIST: Forum for rat genetics PHENOME-LIST: Forum for strain characterization and phenotypic data collection Transgenic-List: http://lists.man.ac.uk/mailman/listinfo/transgenic-list

Transgenic-List is an e-mail forum focusing on issues surrounding the generation, care, and use of genetically engineered mice.

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form the Mouse Mutant Resource (MMR) in 1983. The primary functions of the MMR are to identify and characterize new mutations, to disseminate information through publications and the Mouse Genome Database, and to propagate and distribute these valuable models to the research community. The MMR holds the world’s largest collection of spontaneous mouse mutants. A large number of these were identified from the production colonies by animal-care technicians trained to recognize deviant appearance and behavior. A search of the JAX® Mice database reveals more than 80 different strains that have defects in the eye. Research involving many of these mice is supported by agencies that fund their identification, characterization, and distribution. For example, the Eye Mutant Resource at The Jackson Laboratory is funded by National Eye Institute of the National Institutes of Health and the Foundation Fighting Blindness. Although spontaneous mutations continue to provide a wide variety of models for human disease, their incidence is rare and large-scale screening for desired traits that are not easily observed or measured is time-consuming and not cost-effective. Advances in genetic engineering and phenotyping technologies have made the production and characterization of new models much easier. Genetically engineered mice comprise those created through transgenesis, targeted mutagenesis using homologous recombination, and random mutagenesis. Transgenic mice have defined genetic material randomly added to their genomes.6 Such strains have been used to study gene function and expression and as a result have become important disease models. Strains created by targeted mutagenesis employ homologous recombination to alter or replace a specific locus or gene.7,8 Currently, the majority of strains created by gene targeting carry a null mutation for the gene in question. More recently, conditional targeted mutations have been created that allow control of both the tissue specificity of the mutation and onset of gene expression (temporal control).9,10 Gene targeting produces strains used to study gene function and to create models for human genetic diseases for which the offending gene is known. Random mutagenesis protocols such as treating mouse gametes or ES cells with chemical mutagens11 and gene trapping with retroviral vectors12 are also used to produce valuable new models. These random approaches to mutagenesis produce both dominant and recessive mutations, although most efforts to date have concentrated on identifying the more easily identified dominant mutations. To obtain maximum value from random mutagenesis approaches, rapid and systematized protocols for phenotypic screening must be available, as well as sufficient resources for mapping and cloning genes and subsequently distributing these new models. Several large-scale ENU mutagenesis projects are currently under way.13,14 In addition, governmental agencies are currently funding ENU mutagenesis initiatives in a wide variety of disciplines.15 One of these includes Neuroscience Mutagenesis Facility (JAX-NMF) at the The Jackson Laboratory. The MRC, German Human Genome Project, and JAX-NMF ENU mutagenesis programs are all currently screening ENU-induced mutant mice for vision impairment or defects in eye development. Genetically engineered mice are available from a variety of mouse vendors and special repositories (Table 15.4). The Jackson Laboratory established the Induced Mutant Resource (IMR) in 1993. As the first repository for genetically engineered mouse models, the IMR has served as the prototype for the establishment of newer repositories like the European Mouse Mutant Archive (EMMA) and the Mouse Mutant Regional Resource Centers (MMRRC). These government and privately funded repositories have many functions, including the identification and selection of important mouse models, their importation, cryopreservation, and distribution.

FUTURE DIRECTIONS Genetically defined mice carrying both spontaneous and genetically engineered mutations will continue to be vital tools for increasing our knowledge of the both normal biological processes and diseases of the eye. The study of mouse models, in conjunction with the sequencing of the human

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TABLE 15.4 Genetic Resources for Mice Carrying Induced Mutations Charles River Laboratories, Inc.: http://www.criver.com

Charles River is a commercial distributor of outbred and inbred mice, rats, guinea pigs, hamsters, gerbils, and swine for the biomedical research community.

European Mouse Mutant Archive: (EMMA) http://www.emma.rm.cnr.it

EMMA is a nonprofit repository for transgenic mouse strains essential for basic biomedical research and models for research into complex diseases. These strains can be obtained by qualified research scientists. EMMA consists of several nodes in different locations in European countries; the main site is located in Monterotondo (Rome, Italy).

Harlan Sprague Dawley: http://www.harlan.com

Harlan Sprague Dawley is a commercial distributor of outbred and inbred mice, congenic mice, hybrid mice, mutant mice, and transgenic mice as well as a large number of other laboratory animal stocks and strains. Member of MMRRC.

International Mouse Strain Resources (IMSR): http://www.jax.org/pub-cgi/imsrlist

The IMSR is a multi-center collaboration to promote the use of the mouse as a model organism for research into human diseases, as well as into mammalian physiology and genetics. One of the principal aims of this project is to provide electronically searchable data sets that help the international research community find the necessary resources to carry out fundamental research.

JAX Mice (The Jackson Laboratory): http://www.jax.org/jaxmice

The Jackson Laboratory is a not-for-profit private research institution that distributes the widest range of mice including inbred strains, hybrids, and JAX‚ GEMM strains (Genetically Engineered and Mutant Mice). Resources include federally and privately funded repositories like the Mouse Mutant Resource (MMR), the Induced Mutant Resource (IMR), Eye Mutant Resource Program, and Mouse Mutant Informatics Coordinating Center (MM-ICC) for the MMRRC.

JAX Mice DNA Resource Catalog: http://www.jax.org/resources/documents/dnares/index.html

The Jackson Laboratory Mouse DNA Resource provides genomic DNAs from most of The Laboratory's diverse genetic stocks. High-molecular-weight DNA is prepared by phenol-chloroform extracting nuclei from the tissues of individual mice. The DNA is suitable for Southern blot analysis and amplification by PCR. Most extractions are from spleen or brain and spleen of pedigreed male mice; however, occasionally DNA is extracted from other tissues and some DNA is preserved from female mice.

The Jackson Laboratory Neuroscience Mutagenesis Facility (JAX-NMF): http://www.jax.org/nmf

JAX-NMF is planned as a progressive scientific resource that each year will provide 50 novel murine models useful for the study of important human neurological disorders. Valuable models will be identified using an extensive phenotype-driven approach that targets a broad range of recessive neurological phenotypes and characterizes them sufficiently to attract further detailed study. Mice are undergoing vision screening, primarily for retinal degeneration and mutations that lead to progressive vision impairment.

MRC Mammalian Genetics Unit (MGU, The ENU mutagenesis program at the MRC Mammalian Genetics Unit is screenHarwell): ENU mutagenesis program: ing mice for a wide variety of phenotypes including defects in eye development http://www.mgu.har.mrc.ac.uk/mutaand vision. base Mouse Mutant Regional Resource Centers (MMRRC): http://www.mmrrc.org

The MMRRC Program is a national network of regional breeding and distribution nodes. The goals of the MMRRC include (1) distribution of selected mouse models to the scientific community; (2) storage of mouse embryos or gametes by cryopreservation and subsequent recovery of cryopreserved strains; (3) phenotypic characterization (structural and behavioral) of laboratory mice; (4) genetic quality control of laboratory mice (including genetically modified mice) (5) maintenance of an MMRRC mouse resource database. MMRRC members include Harlan Sprague Dawley; University of North Carolina, Chapel Hill; Taconic Farms, Inc.; UC Davis. The MMRRC Distribution Nodes are coordinated through the Mutant Mouse Informatics Coordinating Center (ICC). The ICC is responsible for designing and developing the necessary software and Web site, installing an electronic network linking the MMRRC Distribution Nodes, and maintaining the MMRRC database.

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TABLE 15.4 (continued) Mymouse.org: http://www.mymouse.org

The goal of mymouse.org is to facilitate international collaborative research between phenotype screening centers and the mouse mutagenesis community, and to promote interactions among scientists interested in analyzing the biological mechanisms linking genes, function, and phenotypes. Mymouse.org features (1) a list of screening laboratories with specific details on the assays performed; (2) direct e-mail links to the principal investigator to discuss and arrange for the analysis of new mutant mice; and (3) an interactive database where individuals can enter new mutants or search for a specific mutant mouse model for collaborative analysis.

Oak Ridge National Laboratory ORNL has a collection of several hundred mouse stocks, most of which propagate (ORNL): mutations induced (over a period of several decades) by radiation or chemicals in http://lsd.ornl.gov/htmouse/mmd- various stages of male or female gametogenesis. These mutation may range from single base-pair changes to rearrangements of various sizes, depending on mutagen and main.htm germ-cell stage. Taconic: http://www.taconic.com

Taconic is a commercial distributor of a wide range of mice and rats for biomedical research with new initiatives in Emerging Models Program and is a member of MMRRC.

ENU–Mouse Mutagenesis Screen The German Human Genome Project (DHGP) has established a research center to Project: perform a large ENU mutagenesis screen in the mouse. The research center consists http://www.gsf.de/isg/groups/enu- of a core facility (GSF, Gene Center) and several associated laboratories. The core facility generates mutagenized F1 and G3 mice which are analyzed by the associated mouse.html laboratories. Furthermore, additional screens can be performed by other research groups that are not participants of the German Human Genome Project.

and mouse genomes, has led to exponential growth in our knowledge of gene function, gene interactions, and other functional regions of the genome. Central repositories like the IMR, MMR, and MMRRC provide mice to the research community and ensure the animal health and genetic quality status in a cost-effective manner. These repositories also provide mouse-related services and expertise as well as central databases in which to store information. Large-scale ENU mutagenesis projects and high-throughput gene-targeting programs such as those carried out at Lexicon (http://www.lexicon.com) and Deltagen (http://www.deltagen.com) are taxing current resources. The generation of literally thousands of new mutant mice a year will compromise our ability to adequately characterize, maintain, archive, and distribute these models. Ideally, a researcher would like to have ready access to all new models. In reality, the resources to support the effort required to make such access possible are not currently available, and compromises will have to be made regarding the availability of new strains. Cryopreservation of embryos and sperm is an alternative to the maintenance of live breeding colonies of strains. Although the recovery of animals from the cryopreserved state is relatively more costly and time-consuming than provision from active breeding colonies, the cost to maintain breeding colonies of infrequently used strains would make the price of animals prohibitively expensive if provided on a cost-recovery basis. In the future, technological advances in areas such as cloning may provide alternatives to traditional methods of propagation and maintenance of mouse models.

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REFERENCES 1. Blake, J.A. et al., The Mouse Genome Database (MGD): integration nexus for the laboratory mouse, Nucleic Acids Res., 29:91, 2001. 2. Ringwald, M. et al., The mouse gene expression database, Nucleic Acids Res., 29:98, 2001. 3. Bult, C.J. et al., Web-based access to mouse models of human cancers: the mouse tumor biology (MTB) database, Nucleic Acids Res., 29:95, 2001. 4. The Gene Ontology Consortium, Gene ontology: tool for the unification of biology, Nat. Genet., 25:25, 2000. 5. Bogue, M., Mouse Phenome Database Web Site, The Jackson Laboratory, Bar Harbor, ME, http://www.jax.org/phenome, June 2001. 6. Gordon, J. W. et al., Genetic transformation of mouse embryos by microinjection of purified DNA, Proc. Natl. Acad. Sci. U.S.A., 77:7380, 1980. 7. Smithies, O. et al., Insertion of DNA sequences into the human chromosomal β-globin locus by homologous recombination, Nature, 317:230, 1985. 8. Mansour, S.L. et al., Disruption of the proto-oncogene int-2 in mouse embryo-derived stem cells: a general strategy for targeting mutations to non-selectable genes, Nature, 336:348, 1988. 9. Gu, H. et al., Deletion of a DNA polymerase beta gene segment in T cells using cell type-specific gene targeting, Science, 265:103, 1994. 10. Kistner, A. et al., Doxycycline-mediated quantitative and tissue-specific control of gene expression in transgenic mice, Proc. Natl. Acad. Sci. U.S.A., 93:10933, 1996. 11. Schimenti, J. and Bucan, M., Functional genomics in the mouse: phenotype-based mutagenesis screens, Genome Res., 8:698, 1998. 12. Friedrich, G. and Soriano, P., Insertional mutagenesis by retroviruses and promoter traps in embryonic stem cells, Methods Enzymol., 225:681, 1993. 13. Hrabe de Angelis, M.H. et al., Genome-wide, large-scale production of mutant mice by ENU mutagenesis, Nat. Genet., 25(4):444, 2000. 14. Nolan, P.M. et al., A systematic, genome-wide, phenotype-driven mutagenesis programme for gene function studies in the mouse, Nat. Genet., 25:440, 2000. 15. Schulhof, J., Restocking the ark: supporting the discovery, creation, and maintenance of new animal models, Lab. Animal, 29:25, 2000.

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Index A Acholeplasma spp., 237 Acrolein, 272 Actb gene, 288 Agarose gel electrophoresis, 286–287 Age control selection, 77–78 fecundity decline, 78 glaucoma, 77–78, 141–144 Aging changes cornea, 11 cortex, 336 lids, 6 Albinism, 256, 336 Alcohol central focal corneal opacities, 70 developmental defects, 164 fetal alcohol syndrome, 150, 229 optic nerve hypoplasia, 229 Alexa 488, 280 Alexa 568, 280 Alkaline phosphatase, 281 Alstrom syndrome, 204 Amacrine cells embryogenesis, 49 function, 33, 35 light response, 320 Ammonium hydroxide, sectioning, 267 Anatomy anterior segment/ocular adnexae, 3–21 posterior segment/orbit, 25–41 Angle closure glaucoma, 136, 137 Anhidrotic ectodermal dysplasia, 114 Anophthalmia description, 148, 150, 151 strain background disease, 70 Anterior ciliary vein, 40 “Anterior hyaloid membrane,” 25 Anterior sclera, 13 Anterior segment/ocular adnexae anatomy, 3–21 anterior uvea, 13–19 aqueous drainage system, 17–19 conjunctiva, 6–7 cornea, 8, 9–12 lacrimal system, 8, 9 lens, 19–21, 175, 177 lids, 3–6 sclera, 11 Anterior subcapsular cataract, 178, 179, 180 Anterior uvea, 13–19 Antigens hyalocytes, 26–27

ki-67 antigen, 309–310 proliferating cell nuclear antigen (PCNA), 310 Aqueous drainage system, 17–19 Aqueous humor features, 13, 17 intraocular pressure, 133–134, 136, 137, 139 production, 15 Arabidopsis thaliana, data sets, 347 Arachnoid mater, 36 Araldite, 274 Arrestin, 215 Astrocytes, 36 Autofluorescence, 283–284 Autosomal dominant cone rod dystrophy, 214 Avidin-biotin system, 281 A-wave, 320, 321, 325, 326 A-wave analyses, 330–331

B Backcrosses data analysis, 85 linkage detection, 83, 86, 87 Bardet–Biedl syndrome, 204 Barrett, G., 337 Behcet’s disease, 169 Biomicroscope, 252 Bipolar cells classification, 33 embryogenesis, 49 function/features, 31, 34, 35 “Bladder cell,” cataracts, 177, 178 BLAST, 88 Blepharitis cause, 115 description, 6, 71, 116 mouse strains, 112 Blepharoconjunctivitis development, 70, 71, 122 intraocular pressure, 319 mouse strains, 237 Blood retinal barrier, 36 Blue opsin, 305, 306 Bouin’s solution advantages/disadvantages, 266, 267 immunohistochemistry, 280 necropsy, 258 Bovine serum albumin (BSA), 281 Bowman’s layer, 9 Brain necropsy, 261, 262, 264 Bromodeoxyuridine (BrdU) staining, 310, 311 Bruch’s membrane, 27, 29, 175 Bulbar conjunctiva, 6

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Burset, M., 88 B-wave, 320, 321, 323–325, 326

C Caenorhabditis elegans, data sets, 347 Calibration film, 253 Carter-Dawson, L., 323 Cataracts “bladder cell,” 177, 178 hypermature cataracts, 179 morphology, 177–182 capsular-epithelial, 178 cortical liquefaction, 178–180 general, 177–178 lens extrusion, 180–182 neoplasia, 182 nuclear/cortical, 178 mouse model, 68–69 ocular/systemic disease, 182 overview, 175 strain background disease, 68–69, 151 types, 176, 180 Cell kinetics, 309–311 Central corneal opacities, 70, 116–117 Central retinal artery, formation, 36 Central retinal vein, 37, 40 Checkerboard pattern stimulus, 334, 336–337 Chediak–Higashi syndrome, 70, 164 Choristomas/hyperplasia, 118 Choroid embryogenesis, 49, 50, 52 features, 27 neovascularization, 170, 171, 172, 198 pathology degeneration, 175 developmental defects, 162–167 microphthalmia, 165–167, 196 neovascularization, 170, 171, 172, 198 platelet storage pool disease, 164–165 retinol developmental defects, 162–164 trauma, 169 uveitis, 168–169 postnatal development, 54 Choroidal circulation, 36 Chromatography (DHPLC), 105 Chromosome substitution strains, 85 Cilia location, 3 pathology, 112 postnatal development, 54 Ciliary body embryogenesis, 51 features, 13, 14, 15–17 postnatal development, 53, 55, 56, 58–59 uveitis, 168–169 Ciliary channels, 15 Ciliary muscle description, 16–17 postnatal development, 57, 58–59

Ciliary muscles, 16–17, 57, 58–59 Ciliary processes, 55, 62 Cloning future possibilities, 351 mapped genetic locus, 82 positional cloning, 82 Coat color abnormalities, 165 Colobomas choroid, 171, 231 microphthalmia, 167, 198 mouse mutations, 231 optic nerve, 229–231 retina, 231 retinal pigment epithelium, 220, 231 vitamin A, 162, 187 Computer databases, see also Resources Expressed Sequence Tag (EST), 88 gene identification, 86, 87 Computer image capture devices, 253 Cone cells features, 31, 33 full-field electroretinogram, 322–324, 326, 329 postnatal development, 60 Cone-rod homeobox gene, 214–215 Confocal microscopy living tissue, 306–307, 308–309 multi-dimensional data sets, 302–306 overview, 300 principles, 300–302 utility, 306–307, 308–309 Conjunctiva embryogenesis, 48, 52 features, 6–7 postnatal development, 55 Conjunctivitis, strains, 71 Contrast threshold, VECP, 337 Controls cell kinetics, 311 fixation, 77, 78 immunohistochemistry, 283 Control selection, 77–79 Copper deficiency/Menkes disease, 234 Cornea aging changes, 11 embryogenesis, 48–49, 50–51, 52 features, 8, 9–12 fibrosis, 307 human diseases, 126, 127, 130, 132 keratitis, 11 neovascularization, 11, 117, 125–127, 128 pathology aqueous outflow/intraocular pressure, 133–134 central corneal opacities, 70, 116–117 dystrophic calcification, 124–125 dystrophic mineralization, 124 edema, 127, 130–132, 133, 134, 137 endothelial proliferation, 132, 135 generalized corneal haze, 117–118, 119, 120 glaucoma, 133–136 hyperplasia/choristomas, 118

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355

infectious disease, 116, 123, 124 keratoplasty, 127 neovascularization, 11, 125–127, 128 ocular surface disease, 120, 122, 128 open eyelids, 71, 122, 123, 144 storage diseases, 127–130 stromal hypoplasia, 118–119, 121 trauma, 122, 123 ulcer, 175, 187 postnatal development, 54, 55, 56, 57, 58–59 stromal mineralization, 11 transparency, 9, 11 Cornea guttata, 130 Corneal thickening, 68 Cortical magnification factor (M), 333, 337 Cortical vacuoles, 178, 180 Corynebacterium spp. blepharitis, 71, 112 microphthalmia/anophthalmia, 151 orbital inflammation, 237 Counterstains, 275, 276, 281, 305 Cranial nerve III, 38 Cranial nerve VI, 38 Crossover events, 83–84 Cryopreservation, 351 Cupric sulfate, 284 C-wave, 320

D Darvasi, A., 86 Databases, see Computer databases; Resources Deltagen, 351 Denaturing high-pressure liquid chromatography (DHPLC), 105 Dendritic melanocytes, 13, 41 Denhardt’s solution, 288 Descemet’s membrane abnormal development, 117 anatomy, 9, 10, 11, 17 embryogenesis, 51 glaucoma, 148 injuries, 122 postnatal development, 55, 58–59 thickening, 130 Development abnormalities alcohol, 164 choroid, 162–167 cornea/sclera, 116–120 optic nerve, 228–234 orbit, 235 overview, 45–46 retina, 196–198 vitreous, 182–185 embryogenesis, 46–53 overview, 45–46 postnatal development adult, 62 cornea, 54, 55, 56, 57, 58–59

days 2 to 4, 53, 55 days 6 to 8, 54, 55–57 days 10 to 14, 57, 58–59, 60 days 21 to 56, 57, 61 newborn to day 2, 53–55 structural development E10 to E12, 46–47 E12 to E14, 48–51 E14 to E17, 49, 50, 51 E17 to birth, 51–53 Developmental glaucoma, 136 DHPLC (denaturing high-pressure liquid chromatography), 105 Diabetes, 72, 198 Diabetic retinopathy, 72 Diaminobenzidine (DAB), 278, 279, 281 Dichromatic vision, 31 Diet, intraocular pressure, 319 Diethylpyrocarbonate (DEPC), 268, 285 Differential interference contrast (DIC) image, 302 Digital computer image capture devices, 253 Digital imaging, 308–309 Digoxigenin (DIG), 291, 293 Dilator muscles, 13, 14 Direct sequence analysis, 105 Diseases, see disease names; Pathology; Strain background disease Dissecting photomicroscope, 252 Distichiasis, 112 Drager, U.C., 334 Drosophila, model, 100 Drosophila melanogaster, data sets, 347 Dura muscle attachment, 38 optic nerve, 36 Dystrophic calcification/mineralization, 124–125

E EAE (experimental allergic encephalomyelitis), 235 EAU (experimental autoimmune uveoretinitis), 168, 169 Edema, cornea, 127, 130–132, 133, 134, 137 Edge effects, 29 Electronic bulletin boards, 348 Electronic listservers, 348 Electron microscopy buffers, 272–273 dehydration, 276 embedding, 275, 276 fixation, 272–275 infiltration/embedding, 276 orientation, 272 postfixation, 274–275 protocols, 275–276 solution preparation, 276 tissue trimming, 272, 273–274 Electrophysiological assessments full-field electroretinogram (ERG), 320–333, 340 overview, 320, 340 visual evoked cortical potentials (VECP), 333–340

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Embed 812, 274 Embedding electron microscopy, 275, 276 light microscopy, 267, 268, 269 tissue processing, 267, 268 Embryonic stem (ES) cells, see ES cell mutagenesis EMMA (European Mouse Mutant Archive), 349, 350 Endogenous enzyme activity, 284 Endophthalmitis, 116, 187 Endothelial proliferation, 132, 135 Endothelium, cornea, 9, 10, 11 En grappe end plates, 38 Entropion, 112 ENU action, 95 dosage, 96 mutation rate, 95, 96 projects, 349, 351 screening and, 99, 101, 105 use, 93, 94–95 Enucleation of the eye, 262–264 Eosinophils, conjunctiva, 6 Episcleral vein, 40 Epon, 274 Epon-Araldite resin, 274, 276 Epoxy resins, 274–275 ERG, see Full-field electroretinogram ES cell mutagenesis advantages, 96, 97 genetic screening, 104 rates, 97 use, 93, 349 EST databases (Expressed Sequence Tag), 88 Ethmoidal artery, 40 Ethmoid bone, 38, 39 Ethmoid foramen, 38 European Mouse Mutant Archive (EMMA), 349, 350 Exophthalmos, 237 Experimental allergic encephalomyelitis (EAE), 235 Experimental autoimmune uveitis, 187 Experimental autoimmune uveoretinitis (EAU), 168, 169 Expressed Sequence Tag (EST) databases, 88 External limiting membrane, 31 Extracellular matrix, 48–49 Extraocular muscles, 38, 40, 49, 50, 51 description, 38, 40 embryogenesis, 49, 50, 51 Extraorbital lacrimal gland, 9 Eyelids, see Lids Eye Mutant Resource at The Jackson Laboratory, 349

F Facial nerve (VII), 6 Fecundity decline, age, 78 Fekete’s acid-alcohol-formalin, 267 Festing, M., 347 Fiber-optic illuminators, 252 Fibroblasts, iris, 13 Fixation

Bouin’s solution, 258 controls, 77, 78 electron microscopy, 272–275, 274–275 formaldehyde, 286 formalin, 266–267, 278 glutaraldehyde, 267, 269, 272, 275 immunohistochemistry, 267, 277, 278 in situ hybridization, 290 light microscopy, 266–267, 268 optic nerve, 264 osmium tetroxide, 272, 273, 275 paraformaldehyde, 267, 268, 269, 272, 290 proliferating cell nuclear antigen, 310 Telly’s fixative, 280 Flash systems, 253, 256 Flash VECP, 335–336 Flexner-Wintersteiner rosettes, 173 Fluorescein isothiocyanate (FITC, green), 280 Fluorescence microscopy, 300–301 Fluorescent probes, 302 Fluorophores, 300 Formaldehyde, Northern hybridization, 286 Formalin, 266–267, 278 Foundation Fighting Blindness, 349 Frameshift mutagen, 96 Frontal arteries, 40 Frontal bone, 38, 39 Fuchs’ endothelial dystrophy, 130 Full-field electroretinogram (ERG) accuracy, 324–325 anesthetics, 327 a-wave analyses, 330–331 dark/light adaptation, 329 examples, 325–326 factors affecting, 326–329 mouse body temperature, 328 multifocal electroretinogram (mERG), 332–333 overview, 320–321, 340 pupil size, 328 recording techniques, 321–322, 323, 329–333 rod/cone responses, 322–324, 326 Fundus camera, 252, 254, 256

G Ganglion cell layer description, 35 embryogenesis, 47, 51, 52 glaucoma, 139, 143 light response, 320 postnatal development, 53, 54, 55, 60, 62 Ganglion cell number identifying control, 82, 83, 84–85, 86 visual pathway, 334 Gapd gene, 285, 288 GBASE (Genomic Database of the Mouse), 346 Gel electrophoresis, 83 Gene detection, 87–89 Gene Expression Database (GXD), 347 Gene identification

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Index databases, 86, 87 mutation detection, 88–89 positional candidates, 87 positional cloning, 88 transcripts, 88 verification, 89 web tools, 88 Gene indices, 88 Gene Ontology Consortium, 347–348 Genetic manipulations, cataracts, 68–69 Genetic mapping, 82 Genetic modifier loci, 81 Genetic resources, 348–349, 350–351 Genetic screens chemical mutagens, 105 genome-wide screens dominant mutations, 97, 98 recessive mutations, 97–99 marked balancer chromosome, 103 modifiers/interactive loci, 94–95, 104–105 multiple alleles at single locus, 101 recessive mutation at single locus, 100 recessive mutation in genomic region, 100–104 screening strategies, 97–105 sequence-based analysis, 105 using deletions, 101–102, 104 Geniculocortical afferents, 334 Genome-wide screen dominant mutations, 97, 98 recessive mutations, 97–99 Genomic Database of the Mouse (GBASE), 346 German Human Genome Project, 349 Glands of Zeis, function, 6 Glaucoma age, 77–78, 141–144 anesthesia, 327 anterior segment findings, 136–139 axonal degeneration, 235 causes, 135 classification, 136 glaucoma-like effects age, 141–144 forkhead transcription factors, 144–148 intraocular pressure, 133–134, 136, 137, 139, 143 lamina cribrosa, 139 Muller cells, 34–35 optic nerve, 139, 143 optic nerve dissection, 264 pigment dispersion glaucoma, 175 posterior segment findings, 139 retinal ganglion cell loss, 139, 143 strain susceptibility, 79 trauma, 139 Glial proliferation, glaucoma, 139 Gliosis, 235 Glutamate, removal/role, 34–35 Glutaraldehyde, 267, 269, 272, 275 Goblet cells (conjunctiva) function, 9 location, 6, 7

357 Golgi apparatus features, 15 lens, 19 Graft rejection, 127 Granulomatous dermatitis, 126 Guanidine thiocyanate, 286 Guigo, R., 88 GXD (Gene Expression Database), 347

H Hamartomas, 68, 118 Harderian gland agenesis, 235 dendritic melanocytes, 41 developmental defects, 162 embryogenesis, 48, 49, 50, 51 function/features, 40–41 innervation, 38 necropsy, 259, 261 neoplasms, 240, 241, 242 postnatal development, 54 Harlequin ichthyosis, 113 Head necropsy, 258–261, 262 Hematoxylin, 278, 281, 311 Hematoxylin and Eosin (H&E) stain, 262 Hermansky–Pudlak syndrome, 70, 164 Herpes simplex disease, 169, 187 High-density oligonucleotide microarrays, 105 Historesin, 268 HMDP (Homology Database and Programs), 346 Homer Wright rosettes, 173 Homology Database and Programs (HMDP), 346 Horizontal cells (inner nuclear layer), function/features, 33, 34 Horizontal sine wave gratings, 337–338 Horseradish peroxidase, 281 Human eye, see also disease names; Mouse model choroid, 27 ciliary body, 15 embryogenesis, 46–47, 136 flash VECP, 336 iris fibroblasts, 113 lens, 19 perivascular glia, 35 trabecular meshwork, 17 VECP, 338 vitreous, 25 Human genome, 87, 88, 94, 109, 349, 351 Human sequence contig, 88 Hyalocytes, 26–27 Hyaloid artery embryogenesis, 47, 49, 52 post natal development, 57, 184 Hyaloid vascular system postnatal development, 53, 55, 57, 60, 198 retinal detachment, 216 Hyaluronan, 29 Hydrocephalus, 72 Hypermature cataracts, 179

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Hyperplasia/choristomas, 118 Hyperthyroidism, 237 Hypoplasia (Meibomian gland), 114, 115

I Ichthyosiform diseases, 113 Identifying specimens, 258 Immunohistochemistry antibody choice, 277–278 endogenous enzyme activity, 284 enzyme detection, 278–279, 281, 282 fixation, 267, 277, 278 fluorescence detection, 279–280, 282–283 incubation with antibodies, 281 interpretation, 283–284 overview, 277 protocols, 282–283 staining, 280–282 IMR (Induced Mutant Resource), 345, 349, 350 Indocarbocyanine Cy-2, 280 Indocarbocyanine Cy-3, 280 Induced genetic defects, 67–68 Induced Mutant Resource (IMR), 345, 349, 350 Inferior oblique muscles, 38 Information resources, 346–348 Inner nuclear layer features, 30, 32, 33–35, 36 postnatal development, 53, 55 Inner plexiform layer description, 35 postnatal development, 55 In situ hybridization fixation, 290 overview, 290 probe labeling, 290–292, 293 protocols, 293–296 Intercrosses data analysis, 85 linkage detection, 83, 86 Internal carotid artery, 36 Internal limiting membrane embryogenesis, 52 retina, 35–36 Interstimulus intervals (ISIs), 331 Intraocular pressure anesthesia for measuring, 318 aqueous humor, 133–134, 136, 137, 139 dimensions/volumes, 314 glaucoma, 133–134, 136, 137, 139, 143 measurement procedure, 314–317 measuring, 313–319 ranges, 313 strains, 313–314, 319 stress, 319 In vivo studies confocal microscopy, 306–307 morphometrics, 308–309 IOP, see Intraocular pressure Iridocorneal angle

development, 56 glaucoma, 145, 146, 147 Iris embryogenesis, 51, 52 features, 13 pathology glaucoma, 143, 148 microphthalmia, 196 platelet storage pool disease, 164 uveitis, 168–169 postnatal development, 53, 56, 58–59 Iris/ciliary body junction, 15 Iris clump cells, 13 Iris pigment, postnatal development, 55 Iris pigment epithelium anatomy, 13–15 developmental abnormalities, 196 Iris root, 13 Iris sphincter features, 13, 14 postnatal development, 55 Iris stroma embryogenesis, 50, 51 features, 13, 14 glaucoma, 141 platelet storage pool disease, 165 postnatal development, 55 Iris transillumination, 164 ISIs (interstimulus intervals), 331

J The Jackson Laboratory (TJL) full-field electroretinogram, 325–326 genetic resource, 348–349 Induced Mutant Resource, 345 information resource, 345, 346, 347, 348–349 multifocal electroretinogram, 332–333 spontaneous mutations, 95 JAX Mice database, 349 JAX-NMF ENU mutagenesis programs, 349, 350

K “Karnovsky’s Fix,” 272 Keeler, C.E., 201 Keratinocytes conjunctiva, 6 cornea, 9 lids, 3 Keratitis, cornea, 11 Keratocytes embryogenesis, 51 features, 9, 10 postnatal development, 55, 58–59 Keratolenticular adhesions, 70 Keratoplasty, 127 Ki-67 antigen, 309–310 Knives, sectioning, 268

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359

L Lacrimal artery, 40 Lacrimal bone, 38, 39 Lacrimal drainage system, 4 Lacrimal gland description, 54 inflammation, 236–237 neoplasms, 235, 240 Lacrimal punctae, 4 Lacrimal system, features, 8, 9 Lactobacillus spp., 71, 237 Lamina cribrosa features, 30, 36 glaucoma, 139 Langerhans cells, 11 Lasers, confocal microscopy, 301 Lateral geniculate nucleus (LGN), 334 LaVail, M.M., 323 Lead citrate counterstain, 275, 276 Leber congenital amaurosis, 214 Lens cataracts, 175–182 embryogenesis, 46, 47, 48, 49, 51, 175 features, 19–21, 175, 177 postnatal development, 53–54, 58–59, 60, 62 sectioning, 266, 267, 268 Leukemia, 172 Levator palpebrae muscles, 5 Lexicon, 351 LGN (lateral geniculate nucleus), 334 Lids aging changes, 6 anatomy, 3–6 coronal section, 5 embryogenesis, 48, 49, 50, 51, 52 features, 3–6 necropsy, 258–261 open lids, 71, 122, 123, 144 pathology, 112–113, 114, 115 postnatal development, 53, 54, 55, 58–59 scanning electron microscopy, 4 vascular supply, 6 Ligaments of Lockwood and Whitnall, 6 Light green counterstain, 281 Light microscopy fixation, 266–267 fixation/embedding in situ hybridization, 268 optic nerve collection, 269 optic nerve embedding/sectioning, 269 paraffin sectioning, 267 plastic sectioning, 268 section interpretation, 269, 270–271 section orientation, 269, 270, 271 tissue processing/paraffin embedding, 267 tissue processing/plastic embedding, 268 Likelihood ratio statistic (LRS), 85 Linkage detection computer mapping, 86 mapping basics, 82–84

mapping complex traits, 84–85 mapping data analysis, 85–86 refining, 86–87 Listing of Inbred Strains of Mice and Rats, 347 Listservers, 348 Little, C.C., 228 LOD score statistic, 85 LRS (likelihood ratio statistic), 85 Luminance effect, VECP, 337 Lupus erythematosis ocular pathology, 170, 198 Sjogren’s syndrome, 235–237 uveitis, 168 Lymphocytes choroid, 27 conjunctiva, 6 Lymphocytic periductal inflammation, 237 Lymphoma, 172–173 Lymphoproliferative disorders, 171–173, 239–240

M Macrophages choroid, 27 conjunctiva, 6 Macular degeneration, 171, 218 Manly, K.F., 85 MAPMAKER/EXP, 85 MAPMAKER/QTL, 85 MapManager QT, 85 MapManager QTX, 85 Mapping, linkage detection, 82–86 Mason, W.T., 300 Mast cells choroid, 27 conjunctiva, 6 Harderian gland, 41 Maxilla bone, 38, 39 Medical Research Council (MRC), 95, 349, 350 Meibomian glands coronal section, 5 ductal ectasia, 114, 126 embryogenesis, 51 excessive sebum production, 71 function, 6, 9 hypoplasia, 114, 115 location, 4 necropsy, 259, 260 pathology, 113–115 postnatal development, 58–59 Melanin pigment, glaucoma, 137, 138 Melanoma, uvea, 173 Melanosomes ciliary body, 15 iris stroma, 13, 14 platelet storage pool disease, 164, 166 retina, 29 Meninges, optic nerve, 36, 37 Menkes disease, 234 mERG (multifocal electroretinogram), 332–333

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Messenger RNA, see also In situ hybridization; Northern hybridization hyalocytes, 27 tenascin, 36 Methyl blue counterstain, 281 M factor (cortical magnification factor), 333, 337 MGD (Mouse Genome Database), 347, 349 MGI (Mouse Genome Informatics), 346, 347–348 Michael Festing’s Listing of Inbred Strains of Mice and Rats, 347 Microphthalmia alleles, 196–198, 199 cataracts, 182 description, 148, 150, 151 developmental defect, 162–164, 165, 167 PHPV, 185 strain background disease, 70, 117, 150 Microsatellite markers, 82–83 Microscopy, see Confocal microscopy; Electron microscopy; Light microscopy Microtome chuck, 175, 267 Millipore-type distillation system, 273 Minsky, M., 300 Missense mutations detection, 89 ENU, 94, 95 ES cells, 97 Mitochondria lens, 19 retina, 31–32 Mitotic rates, measurements, 311 MLC (Mouse Locus Catalog), 94, 346 MLDP (Mouse Linkage Database and Programs), 346 MMGR (Mouse Mutant Gene Resource), 348–349 MMRRC (Mutant Mouse Regional Resource Centers), 345, 349, 350, 351 MMSC (Mouse Mutant Stocks Center), 348–349 Modifiers, see QTL/modifiers identification Mollicutes, 237 Moloney murine sarcoma virus, 196 Morphometrics, 307–309 Motion sensitivity, VECP, 337 MouseBLAST, 347 Mouse Genome Database (MGD), 347, 349 Mouse Genome Informatics (MGI), 346, 347–348 Mouse genome project, 87, 94, 109, 347, 349, 351 Mouse Linkage Database and Programs (MLDP), 346 Mouse Locus Catalog (MLC), 94, 346 Mouse model, see also disease names; Photoreceptors, degenerations fetal alcohol syndrome, 150, 229 human cataracts, 68–69 mutagenesis/phenotyping centers, 100, 105–106 overview, 109, 345, 349, 351 retinal degeneration, 202 Mouse Mutant Gene Resource (MMGR), 348–349 Mouse Mutant Regional Resource Centers (MMRRC), 345, 349, 350, 351 Mouse Mutant Resource (MMR), 345, 348–349, 351 Mouse Mutant Stocks Center (MMSC), 348–349

Mouse Phenome Database, 348 Mouse Tumor Biology (MTB) database, 347 MRC (Medical Research Council), 95, 349, 350 mRNA, see Messenger RNA MTB (Mouse Tumor Biology) database, 347 Muller cells embryogenesis, 49 glaucoma, 34–35 inner nuclear layer, 33–35 inner plexiform layer, 35 light response, 320 nerve fiber layer, 35 photoreceptors, 31, 211 Multi-dimensional data sets confocal microscopy, 302–306 stereology, 308–309 Multifocal electroretinogram (mERG), 332–333 Multiple sclerosis, 234, 235 Mus musculus, data sets, 347 MusPCR database, 346 MusProb database, 346 Mutagenesis/genetic screens building mutant resource, 93–95 future, 105–106 mutagenesis, 95–97 overview, 93 screening strategies, 97–105 Mutant resource, 93–95 Mutations detection, 88–89 interpretation, 79 rate, 95, 96 MX-20 specimen radiography device, 258 Mycoplasma spp., 237 Myelin artifact, 262–263 Myoepithelioma, 241, 243

N National Eye Institute, 349 National Institutes of Health, 349 NCL (neuronal ceroid lipofuscinoses), 211–212 Necropsy enucleation of the eye, 262–264 head, 258–261, 262 orbital glands, 259–261 tissue trimming, 262 Negative controls, immunohistochemistry, 283 Neoplasms fibro-osseous proliferation, 241, 243 Harderian gland, 240, 241, 242 lacrimal gland, 240 lymphoproliferative disorders, 171–173, 239–240 myoepithelioma, 241, 243 neurofibromatosis, 239 pigmented neoplasms, 173–175 retinoblastoma, 173, 174 Sjogren’s syndrome, 235 Neovascularization cause, 118, 125–127

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Index

361

choroid, 170, 171, 172, 198 confocal microscopy, 307 cornea, 11, 117, 125–127, 128 subretina, 77–78 Nerve fiber layer features, 30 retina, 35–36 Netherton’s syndrome, 113 N-ethyl-N-nitrosourea, see ENU Neurofibromatosis, 239 Neuronal ceroid lipofuscinoses (NCL), 211–212 Neuroscience Mutagenesis Facility (JAX-NMF), 349, 350 Nictitating membrane location, 8 lubrication, 40 macrophages, 6 necropsy, 261 postnatal development, 53, 57, 58–59 Night blindness, 215, 329 Nissl substance, 35 NIX annotation pipeline, 88 Non-allelic noncomplementation, 94–95 Nonpigmented epithelium (NPE), 15, 16 Nonradioactive probes, 290–292, 293–294 Nonsense mutations detection, 89 ENU, 95 ES cells, 97 Norrie’s disease, 185, 186, 196 Northern blot analysis, 88 Northern hybridization overview, 285 protocols, 286–290 ribonucleases, 285 total RNA preparation, 286 Nuclear fast red counterstain, 281 Null allele (QTL), gene verification, 89 Nusinowitz, S., 332

O Oak Ridge National Laboratory (ORNL), 95, 351 OCT compound, 280 Ocular adnexae/anterior segment anatomy, 3–21 Ocular development, see Development Ocularity, VECP, 337 Ocular surface disease, 120, 122, 128 Oculomotor nerve (III), 6 Oguchi’s disease (night blindness), 215 Oligodendrocytes, 36 Oligonucleotide primers, 87, 88 Olson, J.M., 85 Open angle glaucoma, 136 Open eyelids, 71, 122, 123, 144 OPs (oscillatory potentials), 320, 321 Opthalmic artery, 36, 40 Opthalmic vein, 36 Optic canal, 38 Optic cup

embryogenesis, 46, 51 features, 13 retina development, 27 Optic nerve brain dissection, 264 dissection, 263 embryogenesis, 47, 48, 50 features, 30, 36–38 glaucoma, 139, 143, 264 light microscopy, 269 pathology aplasia, 228–229 axonal degeneration, 235 coloboma, 229–231 copper deficiency/Menkes disease, 234 cyclopia, 228 degenerations/inflammation, 234–235 demyelination, 234, 235 developmental abnormalities, 228–234 dysmyelination, 231–233 hypoplasia, 229, 230 optic neuritis, 235 overview, 227 Wallerian degeneration, 234 postnatal development, 55, 60 pulling, 263 Ora serrata description, 27 embryogenesis, 53 postnatal development, 57 Orbicularis oculi muscle, 6 Orbit anatomy, 38, 39 pathology agenesis of Harderian glands, 235 developmental abnormalities, 235 lacrimal gland, 235–237 orbital inflammation, 237, 239 Sjorgren’s syndrome, 235–237 thyroid orbitopathy, 237, 238 Orbital cellulitis, 70 Orbital glands location, 9 necropsy, 259–261 Orbital muscles, 6, 38 Orbital venous drainage, 36, 39–40 Orbital venous sinus function, 40 humans, 27 optic nerve dissection, 263 Organ of Corti, 31 Orientation electron microscopy, 272 light microscopy, 269, 270, 271 Oscillatory potentials (OPs), 320, 321 Osmium tetroxide, 272, 273, 275 Osteopetrosis, 165, 196 Outer nuclear layer features, 30, 32 photoreceptor degeneration, 211

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postnatal development, 53 Outer plexiform layer immunohistochemistry, 278, 279 postnatal development, 55, 57

P Palatine bone, 38, 39 Palpebrae, see Lids Panophthalmitis, 187 Paraffin sections immunohistochemistry, 282 in situ hybridization, 293 light microscopy, 267 Paraformaldehyde electron microscopy fixative, 272 in situ hybridization, 290 light microscopy fixative, 267, 268, 269 Parsons, J.H., 45 Pars plana description, 14, 27 development, 57, 62 Pasteurella pneumotropica, 71, 237 Pathology, see also Development, abnormalities; disease names; Strain background disease anophthalmia description, 148, 150, 151 strain background disease, 70 anterior segment, 111–151 choroid, 162–171, 172, 175 cilia, 112 cornea, 116–136, 137 corneal infectious disease, 116, 123, 124 glaucoma, 133–148 iris, 164, 168–169, 196 lens, 175–182 lids, 112–113, 114, 115 Meibomian glands, 113–115 microphthalmia alleles, 196–198, 199 cataracts, 182 description, 148, 150, 151 developmental defect, 162–164, 165, 167 PHPV, 185 strain background disease, 70, 117, 150 neoplasms, 171–175, 239–243 optic nerve, 227–235 orbit, 235–238, 239 persistent hyperplastic primary vitreous (PHPV), 184–187 retina, 195–221 tarsal glands, 115–116 trauma, cornea, 122, 123, 169 uveitis, 168–169 vascular disease, 169–171, 172 vitreous, 182–187, 198, 216 Pattern VECP, 336–338 PBS (phosphate-buffered saline), 280, 282, 283 PCNA (proliferating cell nuclear antigen), 310 PCR database, 346

Pelizaeus–Merzbacher disease (PMD), 231 Pemphigus vulgaris, 113 Pericytes, 36 Perivasculitis, 235 Permutation testing, 85 Peromyscus colony, 325–326 Persistent hyperplastic primary vitreous (PHPV) developmental abnormality, 184–185 ectopic gene expression, 185 Norrie's disease, 185, 186, 196 p53-Null mice, 185 retinoic acid/receptors, 187 Phagosomes features, 29 microphthalmia, 198 Phosphate buffer, 276 Phosphate-buffered saline (PBS), 280, 282, 283 Photodissecting microscope, 252 Photography external ocular photography, 252–254 fluorescein angiography, 256–257 fundus, 256 gross photography, 251–252 overview, 251 slit lamp, 252–253 techniques, 254–255 Photoreceptors degenerations, 198–216 arrestin/Oguchi’s disease, 215 cone-rod homeobox gene, 214–215 degeneration-1, 201, 203 degeneration-2, 201, 203–204 degeneration-3, 204, 205 degeneration-4, 204, 205 degeneration-5, 204–207 degeneration-6, 207, 208 degeneration-7, 207, 209 nervous mutant mouse, 211, 212–213 neuronal ceroid lipofuscinoses (NCL), 211–212 overview, 198–201 Purkinje cell degeneration, 208, 210, 211 rhodopsin mutations, 216 development, 55, 57, 60, 307 embryogenesis, 49 features, 31 PHPV, see Persistent hyperplastic primary vitreous Phthisis bulbi, 169, 170 Physical contig, 88 Pia mater, 36 Pigmented epithelium (PE) ciliary body, 15, 16 photoreceptors, 32 Pineal gland, 31 Pinocytotic vesicles, 15 Pituitary gland, necropsy, 262 Plastic sectioning, light microscopy, 268 Platelet-derived growth factor-A (PDGF-A), 68 Platelet storage pool deficiency, 70 Platelet storage pool diseases, 164–165, 166, 220

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Index

363

Polygenic inheritance, 81–82 Polymorphic markers, 84 Porphyrin, Harderian gland, 41, 48 Positional candidates, gene identification, 86, 87 Positional cloning, 82, 86, 88 Positive controls, immunohistochemistry, 283 Posterior ciliary vein, 40 Posterior segment/orbit anatomy, 25–41 Posterior uvea, 27 Postfixation, electron microscopy, 274–275 Premelanosomes, retina, 29 Primary ocular lymphoma, 173 Primate eye Bruch's membrane, 27, 29 ciliary muscle, 16 cornea, 11 ganglion cells, 334 periorbita, 6 tear fluid, 9 trabecular meshwork, 11, 17 vitreous, 25 Probes fluorescent probes, 302 labeling in situ hybridization, 290–292, 293 nonradioactive probes, 290–292, 293–294 radioactive probes, 290–292, 293 Probes database, 346 Proliferating cell nuclear antigen (PCNA), 310 Propionibacterium acnes, humans, 71 Protein synthesis, photoreceptors, 31 Pulse labeling, 309 Pupillary diameter, control, 13 Purkinje cell degeneration, 208, 210, 211

Q QTL Carthographer, 85 QTL/modifiers identification, 81–89 gene identification, 87–89 linkage detection, 82–87 overview, 81–82, 89 Quantitative morphometrics, 308–309 Quantitative trait locus, see QTL/modifiers identification

R RACE reactions (Rapid Amplification of cDNA Ends), 88 Radioactive labeling, cell kinetics, 311 Radioactive probes, 290–292, 293 Radiographic techniques, 258, 259 RAR (retinoid receptors) null mutants, 162, 163 Rat Data resource, 347 Rd3 retinal degeneration gene, 324–325 Recombinant congenic strains complex trait mapping, 85, 86–87 mapping pitfalls, 87 Recombinant inbred strains background disease, 72 complex trait mapping, 84–85, 86 Rectus muscles, 38

Reeler mouse, 325, 326 Reelin, 325 Resources future, 349, 351 genetic, 348–349, 350–351 information, 346–348 overview, 345 Retina, see also Ganglion cell layer; Photoreceptors Bruch’s membrane, 27, 29, 175 degeneration, 69–70 embryogenesis, 46, 47, 48, 51, 53, 69 features, 26, 27–36 inner nuclear layer, 30, 32, 33–35, 36, 53, 55 inner plexiform layer, 35, 55 nerve fiber layer, 35–36 organization/function, 27, 29 outer plexiform layer, 31–33 pars plana, 14, 27 pathology, 195–221 detachment, 198, 216–217, 221 developmental abnormalities, 162–163, 196–198 microphthalmia, 196–198, 199 neovascularization, 77–78 overview, 195–196 photoreceptor degenerations, 198–216 retinal dysplasia, 46, 185, 196, 197 retinopathy of prematurity, 198, 200 trauma, 169 vascular disease, 198 postnatal development, 54, 55, 57, 60, 62, 69 synapses, 31–33 vascular system, 36, 37 Retinal arterioles, formation, 36 Retinal astrocytic hamartomas, 68 Retinal degenerations identifying control, 82 strain background disease, 69–70 Retinal pigment epithelium (RPE) embryogenesis, 46 features, 28, 29 immunohistochemistry, 283 pathology degenerative/reactive changes, 220–221 dominant negative FGFR–1, 218, 219 gyrate atrophy, 175, 218 hyperplasia, 221 photoreceptor degenerations, 204, 207, 210, 211, 216 senescence-accelerated mouse, 218 vascular disease, 170 photoreceptors, 32 pigmented neoplasms, 173 platelet storage pool disease, 164, 166 retinal detachment, 216 vascular system, 36 Retinal vascular disease, 198 Retinitis pigmentosa, 82, 207, 214, 216, 331 Retinoblastoma, 173, 174 Retinoic acid

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development effects, 162–164 persistent hyperplastic primary vitreous (PHPV), 187 Retinopathy of prematurity, 198, 200 Retinoschisis ERG, 326 Norrie’s disease, 185 Retractor bulbi muscle, 38 Retrolental fibroplasia, 198, 200 Retroviral vectors, 349 Reynolds lead citrate, 275, 276 Rheumatoid arthritis, Sjogren’s syndrome, 235 Rhodopsin kinase, 215 Ribonucleases, Northern hybridization, 285 RI strains, see Recombinant inbred strains Rod cells full-field electroretinogram, 320, 322–324, 326, 329, 330, 331 function, 31, 33 multifocal electroretinogram, 332 postnatal development, 60 “Rodless retina,” see Photoreceptors, degenerations Russell, W.L., 70

S Saccharomyces cerevisiae, data sets, 347 Sample size ENU mutagenesis, 96 linkage detection, 86 Schlemm’s canal anatomy, 17 development, 56, 57, 58–59, 61, 62 glaucoma, 136, 148–149 Schnyder’s crystalline corneal dystrophy (SCCD), 130 Schwalbe’s line mice, 17 primates, 11 Sclera, see also Cornea, pathology features, 11 Scleral canal, 36 Scleroderma, Sjogren’s syndrome, 235 S-cone syndrome, 207 Scotopic threshold response (STR), 320–321 Sebaceous gland ductal ectasia, 126 Sectioning ammonium hydroxide, 267 immunohistochemistry, 282 in situ hybridization, 293 knives, 268 lens, 266, 267, 268 light microscopy, 267, 268 optic nerve, 269 Section interpretation, 269, 270–271 Section orientation, 269, 270, 271 Selection of controls, 77–79 Sex, control selection, 78 Sexual maturity, age, 78 Sickle cell anemia, 169–170 Simple morphometrics, 307 Simple sequence repeat (SSR) marker, 82

Single nucleotide polymorphisms (SNPs), 86 Single-strand conformation polymorphism (SSCP), 105 Sjogren’s syndrome, 235–237 Sjostrand, F.S., 33 Skeletal preparations, 39, 257 Skull necropsy, 261, 262 skeletal preparations, 39, 257 Slit lamp cataract monitoring, 176, 178 description, 252–253 function, 254–255 Slowfade Light Antifade solution, 282 SLT (specific locus test), 95, 96 Small animal fundus camera, 252, 254, 256 SNPs (single nucleotide polymorphisms), 86 Southern analysis, 88 “Spaces of Fontana,” 17 Specific locus test (SLT), 95, 96 Specimen identification, 258 Speed congenic construction, 87 Sphenoid bone, 38, 39 Spiroplasma spp., 237 Splicing alterations, ENU, 95 Spurrs, 274 Squamous cell carcinoma, 113 SSCP (single-strand conformation polymorphism), 105 SSR (simple sequence repeat) marker, 82 Staphylococcus epidermidis, 71 Statistical geneticist, 85 Stereology, 308–309 Stereotaxic apparatus, 335, 337 Storage diseases, 127–130 Strain background disease blepharitis, 71 cataracts, 68–69, 151 conjunctivitis, 71 induced genetic defects, 67–68 microphthalmia, 70, 117, 150 open eyelids, 71, 144 overview, 67–68 platelet storage pool deficiency, 70 recombinant inbred (RI) strains, 72 retinal degeneration-1, 201 retinal degenerations, 69–70 systematic disease, 72 Strains age-related glaucoma, 141–144 age variations, 77–78 control source, 78 disease susceptibility, 79 intraocular pressure, 313–314, 319 Michael Festing’s Listing of Inbred Strains of Mice and Rats, 347 nerve fiber variability, 37 Stress, intraocular pressure, 319 Striated levator muscle, 6, 40 Stroma, cornea, 9, 10 Stromal hypoplasia, 118–119, 121 Stromal mineralization, 11

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365

STR (scotopic threshold response), 320–321 Subretinal neovascularization, 77–78 Sudan Black B, 284 Superior oblique muscles, 38 Superior rectus muscle, 6 Sweep VECP, 337–338 Sympathetic ophthalmia, 169 Synapses, retina, 31–33 Synaptic ribbon, 31–33, 35 Synechias, 137, 143

T Tarsal conjunctiva, 6 Tarsal glands, 115–116 Tarsal (Muller’s) muscle, 6 Tarsal plate, 6 Telly’s fixative, 280 TEM, see Electron microscopy Temporal bone, 38, 39 Temporal tuning function, VECP, 337 Tenascin, 36–37 TEP (transepithelial potential), 320 Tetramethyl rhodamine (TRITC, red), 280 3D data sets, 302–306, 308–309 Thyroiditis, 237 TIGR Gene Index, 88 Tissue processing immunohistochemistry, 280–281 paraffin embedding, 267 plastic embedding, 268 Tissue trimming electron microscopy, 272, 273–274 method, 262 Toluidine blue counterstain, 281 Trabecular meshwork development, 45 embryogenesis, 51 features, 17–19 glaucoma, 136, 137–138, 148–149 postnatal development, 53, 54, 55, 57, 58–59, 61, 62 Transepithelial potential (TEP), 320 Transgenic whole large insert clone rescue, 87, 88 Transmission electron microscopy, see Electron microscopy Trauma choroid, 169 cornea, 11, 122, 123 Trigeminal (V) nerve, 6 Tri-Reagent, 286 Tritiated thymidine, 310, 311 Troland (td), 328 Tub gene, 288, 291 Tub mutant, photoreceptor degeneration, 204–205, 207 Tulp1 gene, 288, 291 Tumor detection, 309, 310 Tunis vasculosa embryogenesis, 51 postnatal development, 55, 57, 62

U Ulceration cornea, 175, 187 lids, 112 Ulcerative blepharitis age, 6 mice strains, 112, 113 open lids, 71 Ultradian rhythms, 336 UNIGENE, 88 Uranyl acetate stain, 276 Ush2a gene, 325 Usher syndrome, 204 Uveal melanoma, 173 Uveitis, 168–169

V Vannas-style scissors, 273, 274 Vascular disease, 169–171, 172 Vascular structures photography, 256 Vascular supply lids, 6 orbit, 39–40 Vascular system, retina, 36, 37 VECP, see Visual evoked cortical potentials Vectashield Mounting Medium, 282 Vestibular organ, ribbon synapses, 31 Visual acuity, 337 Visual cascade, 29 Visual cortex, edge effects, 29 Visual evoked cortical potentials (VECP) mouse visual pathway, 334 multifocal, 338–339 overview, 333, 340 stimuli, 334–338 flash response, 335–336 pattern response, 336–338 Visual pathway, 334 Vitamin A central focal corneal opacities, 70 developmental defects, 162–163 PHPV, 187 Vitreous features, 25–27 function, 15 pathology developmental abnormalities, 182–185 hemorrhage, 187 inflammation, 187, 216 persistent hyperplastic primary vitreous (PHPV), 184–187, 216 vitreous agenesis, 182–183 postnatal development, 60 Vitreous cavity, 46, 47, 51 Vogt-Koyanagi-Harada syndrome, 169 Vortex veins, 40

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W Waardenburg syndrome, 167 Wagor, E., 333, 334 Weaning age, 78 Web site, JAX mouse product list, 201, 347, 350 Web sites cells, 94 gene indices, 88 general information, 346–348 genetic resources, 350–351 mutagenesis/phenotyping centers, 100 recessive mutation screening, 100 Web tools, gene identification, 88 Western blotting, 278 Woolsey, T.A., 336

X X-linked human diseases, 234 X-rays, 258, 259 Xy imaging, 305–306 Xz imaging, 305–306

Z Zygomatic bone, 38, 39

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