Visual Transduction And Non-Visual Light Perception (Ophthalmology Research)

Visual Transduction and Non-Visual Light Perception

Ophthalmology Research Joyce Tombran-Tink, PhD, and Colin J. Barnstable, DPhil SERIES EDITORS

Visual Transduction and Non-Visual Light Perception, edited by Joyce Tombran-Tink, Phd, and Colin J. Barnstable, D Phil, 2008 Mechanisms of the Glaucomas: Disease Processes and Therapeutic Modalities, edited by M. Bruce Shields, MD, Joyce Tombran-Tink, PhD, and Colin Barnstable, DPhil, 2008 Ocular Transporters in Ophthalmic Diseases and Drug Delivery, edited by Joyce TombranTink, PhD, and Colin J. Barnstable, DPhil, 2008 Visual Prosthesis and Ophthalmic Devices: New Hope in Sight, edited by Joseph F. Rizzo, MD, Joyce Tombran-Tink, PhD, and Colin J. Barnstable, DPhil, 2007 Retinal Degenerations: Biology, Diagnostics, and Therapeutics, edited by Joyce TombranTink, PhD, and Colin J. Barnstable, DPhil, 2007 Ocular Angiogenesis: Diseases, Mechanisms, and Therapeutics, edited by Joyce TombranTink, PhD, and Colin J. Barnstable, DPhil, 2006

Visual Transduction and Non-Visual Light Perception Edited by Joyce Tombran-Tink, PhD Department of Ophthalmology Department of Neural and Behavioral Sciences Milton S. Hershey Medical Center Penn State University College of Medicine, Hershey, PA, USA

Colin J. Barnstable, DPhil Department of Neural and Behavioral Sciences Milton S. Hershey Medical Center Penn State University College of Medicine, Hershey, PA, USA

Editors and Series Editors Joyce Tombran-Tink, PhD Department of Ophthalmology Department of Neural and Behavioral Sciences Milton S. Hershey Medical Center Penn State University College of Medicine Hershey, PA, USA

Colin J. Barnstable, DPhil Department of Neural and Behavioral Sciences Milton S. Hershey Medical Center Penn State University College of Medicine Hershey, PA, USA

ISBN: 978-1-58829-957-4 e-ISBN: 978-1-59745-374-5 DOI: 10.1007/978-1-59745-374-5 Library of Congress Control Number: 2008925918 © 2008 Humana Press, a part of Springer Science + Business Media, LLC All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, 999 Riverview Drive, Suite 208, Totowa, NJ 07512 USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Cover illustration: Figure 2, Chapter 1, “An Organ of Exquisite Perfection,” by George Ayoub. Adapted from a diagram from www.webvision.med.utah.edu. Modified by Nancy Fallatt. Back cover images from Figure 1, Chapter 17, “Multifocal Oscillatory Potentials of the Human Retina,” by Anne Kurtenbach and Herbert Jägle. Printed on acid-free paper 9 8 7 6 5 4 3 2 1 springer.com

Preface Sensitivity to light is a near-universal attribute of living organisms. It can be seen in the tropic responses of plants, in movements of many bacteria and unicellular organisms, as well as in the more complex visual responses of most animals. While the mechanisms of light detection and the behavioral consequences of its detection in these species are a series of fascinating stories, this volume is concerned with vision in its more classical interpretation. Although ancient philosophers, including Aristotle and Ptolemy, thought that the function of the eye was to emit light and illuminate objects, it has been over a thousand years since the Persian Alhazan (Abu Ali Hasan Ibn al-Haitham) explained that vision was the result of light coming from an object into the eyes. What happened to the light after traversing the optical path of the eyes remained unclear for many centuries. Leonardo daVinci and others of that era thought that light was channeled back to the ventricles of the brain through the optic nerves. In the early nineteenth century, as the structure of the eye, and particularly the retina, were examined more carefully, it became apparent that vision was linked to a transformation process, which occurred in the retina and specifically in the photoreceptors. The visual pigments and their sensitivity to light were described in the mid-nineteenth century. In the first half of the twentieth century, the pioneering work of Wald showed that the visual pigment was a protein with an attached molecule that had properties of a carotenoid. The identification of this chromophore and its derivatives as retinal and retinol and the enzymatic conversion of one to the other were landmark studies carried out in the laboratories of Morton in England and Wald in the United States. This finding was the cornerstone for the next major breakthrough in our understanding of the visual transduction cascade. In a follow-up study, Wald and his coworkers Hubbard and Brown found that the active visual pigment chromophore was 11-cis retinal, and that light induced a transition in this pigment to the all-trans form. We now know that the light-induced change in the conformation of 11-cis retinal is the fundamental step in converting light energy into chemical energy in the retina. The next major breakthrough in our understanding of the visual transduction cascade, the conversion of this cis–trans isomerization of the opsin chromophore, part of the rhodopsin complex, into changes in membrane conductance and synaptic signaling took another 30 years to understand. The work during this period showed that the lightsensitive rhodopsin machinery is primarily located in disks that are completely isolated from the plasma membrane but electrical signals involve changes in conductance at the plasma membrane of the rod photoreceptors. Perhaps the most important realization in this story is that rod photoreceptors need an internal signal molecule. For many years, the two rival candidates for this internal signal were calcium and cyclic guanosine monophosphate (cGMP). Physiological measurements showed that changes occurred in calcium fluxes in rod photoreceptors on illumination, a finding that later led to the identification of a biochemically defined light-sensitive enzymatic machinery that hydrolyzed cGMP. The critical role of cGMP v

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in visual transduction was later established when Fesenko showed that this nucleotide could directly regulate the opening of a novel class of membrane channels, the cyclic nucleotide-gated cation channels. Thus, the pathway from light to an alteration in rod photoreceptor membrane conductance was finally established in the twentieth century and over the past decade has been characterized in much greater detail. We now know that photoreceptors cannot regenerate 11-cis retinal by themselves, and that all-trans retinal from the photoreceptors is carried via a number of retinoid-binding proteins to the retinal pigment epithelium (RPE) cells, where it is regenerated to 11-cis retinal. We also know that photoreceptors have the remarkable ability to adapt to different levels of background illumination with minimal loss of sensitivity, and that visual transduction by itself is not sufficient to create signals that can be transmitted back to the visual cortex. The information from photoreceptors passes through many types of retinal neurons, and a highly processed signal is sent back to visual centers in the brain through the ganglion cell axons so that the signals can be interpreted. Thus, we have found that vision is a much more complex and dynamic process than those initially proposed by the ancient philosophers, and that it occurs through an exquisite biochemical transduction system made possible through the concerted effort of all cell types in the retina. In this text, the authors discuss many important facets of the visual transduction cascade, including photoreceptor membrane conductance, how the RPE regenerates 11-cis retinal, photoreceptor adaptation to various levels of illumination and the biochemical basis of this phenomenon as well as its psychophysical consequences, how the retina develops into its final structure, how signals are processed in the retinal synaptic layers, and how changes in the retina and RPE influence normal aging. An important message in this volume is that as we continue to understand the molecular and biochemical intricacies of visual transduction and the many aspects of aging and retinal degeneration, we can adopt a series of dietary and lifestyle changes and with pharmaceutical aids can slow the decline in visual function. Whether this will be enough to stave off loss of vision or onset of age-related disease remains to be seen. Loss of vision is paralyzing to individuals, their family members, and the health care system. The recent statistics from the National Eye Institute show that there is an increase in the numbers of the elderly with visual impairment, and that this will continue to rise with the burgeoning aging population. Thus, there is an urgent need to understand the biochemical mechanisms that allow us to see and to study how these mechanisms are affected by aging and pathology so that better therapeutics can be developed to make vision possible at all stages of our lives. Joyce Tombran-Tink Colin J. Barnstable

Contents Preface..................................................................................................................... Contributors ............................................................................................................ Companion CD ....................................................................................................... Part I

Evolution of the Visual System

1 An Organ of Exquisite Perfection ..................................................... George Ayoub Part II

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3

Photoreceptor Structure, Function, and Development

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Development of the Foveal Specialization ....................................... Keely M. Bumsted O’Brien

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An Update on the Regulation of Rod Photoreceptor Development .. Edward M. Levine and Sabine Fuhrmann

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Part III 4

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The Retinal Pigment Epithelium and the Visual Cycle

Photoreceptor–RPE Interactions: Physiology and Molecular Mechanisms .......................................................... Silvia C. Finnemann and Yongen Chang Molecular Biology of IRBP and Its Role in the Visual Cycle .......... Diane E. Borst, Jeffrey H. Boatright, and John M. Nickerson

Part IV

Regulation of Photoresponses by Phosphorylation .......................... Alecia K. Gross, Qiong Wang, and Theodore G. Wensel

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The cGMP Signaling Pathway in Retinal Photoreceptors and the Central Role of Photoreceptor Phosphodiesterase (PDE6)............................................................ Rick H. Cote

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Visual Signaling in the Outer Retina

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67

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Rhodopsin Structure, Function, and Involvement in Retinitis Pigmentosa ................................................................. Scott Gleim and John Hwa

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Multiple Signaling Pathways Govern Calcium Homeostasis in Photoreceptor Inner Segments ............................ Tamas Szikra and David Krizaj

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The Transduction Channels of Rod and Cone Photoreceptors ......... Dimitri Tränkner vii

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Rhodopsins in Drosophila Color Vision ........................................... David Jukam, Preet Lidder, and Claude Desplan

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INAD Signaling Complex of Drosophila Photoreceptors ................ Armin Huber and Nina E. Meyer

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Part V 13

Visual Processing in the Inner Retina

Visual Signal Processing in the Inner Retina .................................... Botir T. Sagdullaev, Tomomi Ichinose, Erika D. Eggers, and Peter D. Lukasiewicz

Part VI 14

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Color Vision and Adaptive Processes

Human Cone Spectral Sensitivities and Color Vision Deficiencies ...................................................... Andrew Stockman and Lindsay T. Sharpe

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Luminous Efficiency Functions ........................................................ Lindsay T. Sharpe and Andrew Stockman

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Cone Pigments and Vision in the Mouse .......................................... Gerald H. Jacobs

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Multifocal Oscillatory Potentials of the Human Retina ................... Anne Kurtenbach and Herbert Jägle

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Part VII

Aging and Vision

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The Aging of the Retina ................................................................... Caren Bellmann and José A. Sahel

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Aging of the Retinal Pigment Epithelium ........................................ Michael E. Boulton

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Visual Transduction and Age-Related Changes in Lipofuscin ......... . . Małgorzata Rózanowski and Bartosz Rózanowski

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Part VIII 21

Nonphotoreceptor Light Detection and Circadian Rhythms

A Nonspecific System Provides Nonphotic Information for the Biological Clock ........................................... Marian H. Lewandowski

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22 The Circadian Clock: Physiology, Genes, and Disease .................... Michael C. Antle

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Index .........................................................................................................

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Contributors Michael C. Antle, PhD • Department of Psychology, University of Calgary, Calgary, Canada George Ayoub, PhD • Neuroscience Research Institute and Department of Molecular Cellular and Developmental Biology, University of California, Santa Barbara, CA Colin J. Barnstable, DPhil • Department of Neural and Behavioral Sciences, Milton S. Hershey Medical Center, Penn State University College of Medicine, Hershey, PA Caren Bellman, PhD • Centre Hospitalier National d’Ophtalmogie des Quinze-Vingts, and INSERM U 592, Paris, France Jeffrey H. Boatright, PhD • Department of Ophthalmology, Emory Eye Center, Emory University, Atlanta, GA Diane E. Borst, PhD • Department of Anatomy, Physiology, and Genetics, Uniformed Services University of the Health Sciences, Bethesda, MD Michael E. Boulton, PhD • Department of Ophthalmology and Visual Sciences, University of Texas Medical Branch, Galveston, TX Keely M. Bumsted O’Brien, PhD • Department of Optometry and Vision Science, University of Auckland, Auckland, New Zealand Yongen Chang, PhD • Dyson Vision Research Institute, Weill Medical College, New York, NY Rick H. Cote, PhD • Department of Biochemistry and Molecular Biology, University of New Hampshire, Durham, NH Claude Desplan, PhD • Department of Biology, New York University, New York, NY Erika D. Eggers, PhD • Department of Ophthalmology and Visual Sciences, Washington University School of Medicine, St. Louis, MO Silvia C. Finnemann, PhD • Dyson Vision Research Institute, Weill Medical College, New York, NY Sabine Fuhrmann, PhD • Department of Ophthalmology and Visual Sciences, John A. Moran Eye Center, University of Utah, Salt Lake City, UT Scott Gleim, MS • Department of Pharmacology and Toxicology, Dartmouth Medical School, Hanover, NH Alecia K. Gross, PhD • Department of Vision Sciences, School of Optometry, University of Alabama, Birmingham, AL Armin Huber, PhD • Department of Biosensorics, Institute of Physiology, University of Hohenheim, 70599 Stuttgart, Germany John Hwa, MD, PhD • Department of Pharmacology and Toxicology and of Medicine, Dartmouth Medical School, Hanover, NH Tomomi Ichinose, MD, PhD • Department of Ophthalmology and Visual Sciences, Washington University School of Medicine, St. Louis, MO Gerald H. Jacobs, PhD • Neuroscience Research Institute, University of California, Santa Barbara, CA Herbert Jägle, MD • University Eye Hospital, Tübingen, Germany ix

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David Jukam, BS • Department of Biology, New York University, New York, NY David Krizaj, PhD • Departments of Ophthalmology and Physiology, University of Utah School of Medicine, 65 N Medical Drive, Salt Lake City, Utah, U.S.A. Anne Kurtenbach, PhD • University Eye Hospital, Tübingen, Germany Edward M. Levine, PhD • Department of Ophthalmology and Visual Sciences, John A. Moran Eye Center, University of Utah, Salt Lake City, UT Marian H. Lewandowski, PhD • Department of Neurophysiology and Chronobiology, Institute of Zoology, Jagiellonian University, Kraków, Poland Preet Lidder, PhD • Department of Biology, New York University, New York, NY Peter D. Lukasiewicz, PhD • Department of Ophthalmology and Visual Sciences, Washington University School of Medicine, St. Louis, MO Nina E. Meyer, PhD • Department of Biosensorics, Institute of Physiology, University of Hohenheim, 70599 Stuttgart, Germany John M. Nickerson, PhD • Department of Ophthalmology, Emory Eye Center, Emory University, Atlanta, GA . Małgorzata Rózanowski, PhD • School of Optometry and Vision Sciences, Cardiff University, United Kingdom . Bartosz Rózanowski, PhD • Department of Cell Biology and Genetics, Institute of Biology, Pedagogical Academy of Kraków, Poland Botir T. Sagdullaev, PhD • Department of Ophthalmology and Visual Sciences, Washington University School of Medicine, St. Louis, MO José A. Sahel, MD • Centre Hospitalier National d’Ophtalmogie des Quinze-Vingts, and INSERM U 592, Paris, France Lindsay T. Sharpe, PhD • Institute of Ophthalmology, London, United Kingdom Andrew Stockman, PhD • Institute of Ophthalmology, London, United Kingdom Tamas Szikra, PhD • Friedrich Miescher Institute for Biomedical Research, Maulbeerstrasse 66, 4058 Basel, Switzerland Joyce Tombran-Tink, PhD • Department of Ophthalmology, Department of Neural and Behavioral Sciences, Milton S. Hershey Medical Center, Penn State University College of Medicine, Hershey, PA Dimitri Tränkner, PhD • Institut für Biologische Informationsverarbeitung, Forschungszentrum Jülich, Germany Qiong Wang, PhD • Department of Biochemistry and Molecular Biology, Baylor College of Medicine, Houston, TX Theodore G. Wensel, PhD • Department of Biochemistry and Molecular Biology, Baylor College of Medicine, Houston, TX

COMPANION CD Black and White and Color illustrations are provided on the Companion CD attached to the inside back cover. The image files are organized into folders by chapter number and are viewable in most Web browsers. The CD is compatible with both Mac and PC operating systems.

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1 An Organ of Exquisite Perfection George Ayoub CONTENTS Optical Path Retinal Photoreception Retinal Pathways References

OPTICAL PATH As can be seen in Fig. 1, the eye is a nearly spherical structure, structurally limited by the sclera and cornea. The sclera is the tough white tissue that delimits the outer orbit of the eye, while the cornea is the clear portion in the front. The cornea is a focusing element for the visual path, providing more than half of the focusing power of the eye, with the lens handling the remainder. Focusing of light by the cornea and lens is necessary to create an image on the retina, which is the light-sensitive portion of the eye. This is much like the role of a camera lens in creating a clear image at the CCD (charge-coupled device) or film plane. Indeed, the cornea and lens are the two elements that focus the light, with the cornea fixed in focal length and the lens adjustable. They provide a double positive lens arrangement (i.e., two convex lenses) to accomplish this. Light is focused based on the cornea’s shape and refractive index. The cornea is a nearly spherical structure, slightly flattened to reduce spherical aberration. The index of refraction for light (n) as it passes through the cornea is 1.376. Since the index of refraction in air is 1.0, this change in the refractive index, along with the cornea’s convex shape, causes light rays to bend in a converging manner as they pass through [1–3]. The lens of the eye has a shape that is malleable, allowing this second focusing element in the visual path to be used to adjust the focal point, which allows us to form a clear image on the retina for objects from near to distant. The shape of the lens changes due to the action of two structures. The connective tissue encompassing the lens keeps it spherical, but the suspensory ligaments that surround it pull it into a flatter shape. The ciliary muscles counteract the pull of these ligaments and allow the lens to become more round. Thus, by contracting the ciliary muscles, we are able to focus on objects that are close at hand by increasing the curvature of the lens, thereby increasing its refractive power. Relaxation of this muscle allows us to focus on objects at a distance. From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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Fig. 1. Schematic of the eye and a corresponding micrograph of the retina. The eye structures are labeled and the retinal layers indicated as follows: retinal pigment epithelium (RPE); outer segments of photoreceptors (OS); inner segments of photoreceptors (IS); outer nuclear layer (ONL), which is composed primarily of photoreceptor cell nuclei; outer plexiform layer (OPL), which is the site for synaptic contact between photoreceptors, horizontal cells, and bipolar cells; inner nuclear layer (INL), the location of cell bodies of bipolar cells and most horizontal and amacrine cells; inner plexiform layer (IPL), the site of synaptic contact between bipolar cells, amacrine cells, and ganglion cells; ganglion cell layer (GCL), the location of most ganglion cell somata; and nerve fiber layer (NFL), comprised of the axons of ganglion cells.

As we age, the lens becomes less elastic, and the ability of it to round up decreases. The result is that our nearest clear point of vision moves further away with age. This situation is termed presbyopia [1, 2]. The refractive index of the lens is not a constant. The lens itself is made of over 2,000 layers of cells, with the refractive index of this tissue increasing at the center and being less at the front and back surfaces. The index of refraction at the cortex is 1.386, while the index at the inner core is 1.406. This means that the refractive index is matched to the cornea at the point where light enters the lens and then steps up and down as it passes through. A significant result of this feature is a decrease in reflection of light. As one can observe when light passes through glass (which has an index of refraction of about 1.5), some of the light is reflected. Indeed, this reflection is due to the light passing through the large change in refractive index. As light passes across a large change in refractive index, a portion of the light is reflected, and the larger the difference in the refractive indexes, the greater the reflection that is seen. For clear window glass, this amount of light reflected is 4% of the light at each surface (thus, 92% of the light passes through, with 4% reflected at the front surface of the window and an additional 4% at the back surface of the window). Because the refractive index of the lens changes gradually, less light is reflected as the

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light passes through, allowing for transmission of more of the light and virtual elimination of internal reflections that would otherwise compromise visual acuity [1–3]. Between the cornea and the lens is the iris. This structure provides an aperture that controls the amount of light that enters the eye and passes through to the retina. The iris is located between the two refractive surfaces, reducing distortion of the entering light. The iris is pigmented, and the opening of the iris is termed the pupil. This opening is reduced in brighter illumination and increased in dimmer illumination to provide sufficient stimulation for the retinal cells. Directly behind the cornea is a fluid-filled chamber; the fluid is called the aqueous humor. Fluid is continually secreted into this chamber and continually drained from it. In certain medical conditions, the drain rate of fluid is reduced, and pressure can build. This occurs in many types of glaucoma, in which the increase in pressure is gradual over time and can be controlled to some extent by medication that reduces fluid production. An uncontrolled elevation of pressure causes a gradual loss of vision due to damage to the cells at the back of the eye that form the optic nerve [2]. While the aqueous humor fluid helps maintain the structure and curvature of the cornea, it is the gelatinous vitreous humor that maintains the full shape of the orb of the eye. The vitreous fills the region behind the lens and in front of the retina and provides sufficient pressure to keep the round shape of the eye. Optically, then, light passes through the cornea, aqueous humor, iris, lens, and vitreous humor on its path to the retina. The cornea and lens are the refractive elements, with the lens the only adjustable one, and the iris modulates the amount of light that passes. If there are aberrations in the shape of the eye or of the cornea, then distortion of vision occurs. When the cornea is not radially symmetric, astigmatism results, by which the focus at one angle differs slightly from the focal distance at another angle. If the overall shape of the cornea is rounder or flatter, then the conditions of myopia (nearsightedness) or hyperopia (farsightedness) result because the focal distance is slightly different from what is needed to form a clear image on the retina. Thus, slight changes in the shape of the cornea, since it is the primary focusing element, result in deviations from normal focus. For those who are myopic, the decrease in lens malleability seen in presbyopia has a delayed impact on vision compared to the hyperopic individual. The reason is simply that as the near point of vision moves further from the eyes, if one has started with a closer near point of vision (as is the case in myopia), then the effect is to move the near point to a normal reading distance, while for the normal sighted or hyperopic, it is to move the nearest point of vision to a distance further than is readily readable (indeed, one typically finds that the letters are too small when they are in focus, and so use of reading glasses or a magnifying lens is needed to compensate). RETINAL PHOTORECEPTION Photoreception has two major components: optical and biochemical. The optical component is primarily described in the preceding section, although there are critical retinal portions to this. The biochemical component occurs in the outer retina.

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Fig. 2. The organization of the retina. (Courtesy of Dr. Helga Kolb.)

Photoreception Optics Optically, light focused by the front of the eye is imaged onto the photoreceptor layer of the retina. The retina itself is a thin neural tissue that lines the inside orbit of the eye and is comprised of several layers of cells, as can be seen in Fig. 1. These cells are in repeating arrays, such that in any given area of retina, the same cell types are found in a similar arrangement. The optical arrangement is that light passes first through ganglion cells, then amacrine, bipolar, and horizontal cells before arriving at the photoreceptors, which make up the outer retina (see Fig. 2) [1–4]. The retinal layers are defined from the inner to outer regions of the eye, with the ganglion cell axon fibers the innermost layer of the retina and the photoreceptors and epithelium the outermost retinal layers. Within the retina, there are two types of photoreceptors: The rod photoreceptors provide for our night vision and are exquisitely sensitive to small amounts of light; the cone photoreceptors provide color vision. Cone photoreceptors are responsive to one of three colors (red, green, or blue); combined, the relative intensities of these three primary colors that we can see allow us to determine the color of any object and to see the full spectrum of the rainbow. This feature of only detecting three colors is why we can see all colors when we look at an LCD (liquid crystal display) screen, which is also only comprised of red, green, and blue (RGB), or at a magazine photo, which is typically made from cyan, magenta, and yellow (the complementary colors to RGB) along with black. The central part of the retina, called the fovea, contains a very high number of cone photoreceptors packed into a very small space, about 160,000 cone photoreceptors per square millimeter. This high density of cones is why when we look

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straight at an object we can make out a large amount of detail. The cones in the fovea then provide our highest visual acuity, which is 0.017° (300 µrad, or 1.03 arc minute) of our visual field. This works out to be a 19-inch object a mile away or the size of the pixels on a television or LCD monitor at normal viewing distance. A simple exercise to observe this is to view a computer monitor at a close distance and note that you can see the individual pixels, while at normal distance they are indistinguishable [1, 3]. The fovea provides this high acuity for the central 2° of our visual field. Within this region, there is a pit in the retina as the other cells are angled to the sides of the fovea, eliminating these cells from the path of the light onto the cones. Although the retinal neurons are transparent, for the centralmost vision, the small refraction that the cells create would decrement acuity slightly; thus their angling away from the fovea removes this potential detriment, retaining our crisp vision [1, 3]. Away from the fovea, there is a mixture of rod and cone photoreceptors, and the acuity is much more limited, although the sensitivity to low light levels is higher due to the presence of rods. One can observe this on a dark night, such as when something catches your eye and you turn to look at it, but it is not as bright as when seen in the periphery of one’s vision. Outside of the neural retina lies the retinal pigment epithelium (RPE). This RPE has two essential functions for photoreception, one optical and the other biochemical. Optically, the RPE cells are black, and they surround the outer segments of the photoreceptors. This means that any light that passes around a photoreceptor is absorbed by the RPE. Thus, no light is reflected back onto the retina. This black curtain keeps us from seeing halos around objects and is therefore essential for high acuity. Notably, in nocturnal animals, such as cats, the epithelium is reflective, allowing them to capture every bit of light as they reflect the photons that are not captured in the first pass through. Their vision in dim light is thus very good, but their acuity is not as high as it is for humans due to this trade-off [5, 6]. Photoreception Biochemistry The other key function of the RPE for photoreception is in regeneration of the visual pigment of the photoreceptors. For this, the RPE cells use vitamin A. The RPE exchanges and renews the photosensitive molecule from the photoreceptors, creating an intimate collaboration between these two cells. Hence, the photoreceptors rely on the RPE for both optical purposes and biochemical function. Photoreception takes place in the outer segments of the photoreceptors. Photoreceptors are segmented into three regions: the outer segment, which is packed with photosensitive molecules; the inner segment, which contains the cell nucleus and a large amount of mitochondria to generate the energy for this photoreception; and the synaptic terminal, which communicates to the next cells in the visual system. The biochemical process of photoreception is referred to as phototransduction. Molecules of visual pigment called opsin are embedded within the membrane of the rods and cones. Indeed, the rods and cones have an elaboration of membrane in their outer segments, with the cones having membrane folded over and over on itself and the rods containing a stack of membranous disks, which look much like a large stack of pancakes, within a sleeve of rod membrane. All this membrane is loaded with opsin

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molecules (called rhodopsin in the rods), and these opsins absorb light of a specific range of wavelengths. Rhodopsin absorbs light of about 500 nm, while cone opsins absorb light of 420, 530, and 560 nm (blue, green, and red, respectively). These are the peak wavelengths, meaning that at these colors the opsin molecules are maximally stimulated. They absorb light 50 nm or more greater and lesser than their optimal color, but less strongly, so their response to light decreases as the color is further from their optimal wavelength [1, 4, 7]. When light is absorbed by rhodopsin, the light energy is absorbed by the portion of the molecule called retinal. This retinal is what is regenerated from vitamin A. When retinal absorbs light, its shape changes from a bent form to a straight form (from 11-cis retinal to all-trans retinal). This change in shape begins a cascade of events that result in a change in the voltage across the membrane of the photoreceptor. Briefly, the stimulation of rhodopsin (changing the shape of retinal) results in activation of a molecule called transducin. Transducin is a guanosine triphosphate (GTP)-dependent protein (also called a G protein, part of a family of signaling molecules within many cells). The activation of transducin in turn activates an enzyme called phosphodiesterase. Phosphodiesterase is an enzyme that catalyzes a reaction in which cyclic guanosine monophosphate (cGMP; a signaling molecule within cells) is broken down to GMP. While the cyclic form of GMP is a signaling molecule, the noncyclic form is not. The removal of cGMP from photoreceptors results in a change in the membrane potential of the cells [7]. Photoreceptors have a collection of membrane-spanning channels that allow specific ions to pass through. One of these channels is specific for Na+ ions (sodium, a positively charged ion). This channel requires the presence of cGMP in the cell for its gate to open and Na to travel through. Since cells have more Na outside than inside, when the Na gate is open, the positively charged Na ions enter the cell, making the cell more positive in charge than it had been. These cGMP-gated Na channels then are open in the dark but become closed when the cells see light. Thus, when stimulated by light, photoreceptors become more negatively charged. This change in their membrane voltage is the first electrical sign that we have seen the light and is communicated to the next cells in the retina [1, 4]. Membrane Voltages The voltage across the membrane of a cell is a standard feature of all cells of the nervous system, and it is the change in this membrane voltage that is used by neurons to process and transmit information. Thus, for all neurons, small changes in their membrane voltage are elicited by incoming information, with the incoming information arising from other neurons or from sense organs. These voltage changes (additive or subtractive, as some changes are positive charges and some are negative charges) result in a net change to the cell’s membrane voltage. In certain neurons, when this excursion in the membrane voltage reaches a threshold voltage level, the cell creates a large spike in its membrane voltage called an action potential. The action potential is regenerative, so the electrical spike travels along the thin tubular extension of the neuron called the axon and travels along this axon at a high rate of speed (over 100 m/s in some cells). The amount of communication between neurons is determined in large part by the size of the change in the membrane voltage of the first cell. Now, while the change in

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membrane voltage is variable, the spikes along the axon are all-or-none. To accomplish this, there is a conversion of an analog signal (the gradual changes in membrane voltage) to a digital signal (the pulses or spikes along the axon) that occurs within the neuron. This conversion is very much like the action of an analog-to-digital converter seen in electronics as, for example, when a visual image is converted to a series of digital bits by the CCD of a digital camera. This pulsatile or digital signal is then reconverted back to an excursion in membrane potential (or analog signal) at the synapse. The synapse is a specialized region where two neurons come very close to each other. At the synapse, the first cell (the presynaptic neuron) releases a chemical called a neurotransmitter in amounts proportional to the number of action potentials. The postsynaptic neuron has receptor molecules in its membrane that bind to this neurotransmitter and open gates for ion channels, resulting in a change in the membrane voltage of the postsynaptic cell. Most neurons have many synapses, so all the changes in membrane voltage sum to create the new membrane voltage of the postsynaptic neuron, which determines its action potential rate and amount of neurotransmitter release onto a third neuron. This process continues, with the synapse essentially performing a mathematical function on the information transmitted. The result is that some synapses will simply add signals, some will determine their rate, some will integrate, and so on. In this way, our synapses act as individual processors, with their processing determined both by the chemical used as the neurotransmitter at that synapse and the molecule used as its receptor. These two combine to determine the type of processing that occurs at any one synapse. Given that our brains have about 100 billion neurons, and each neuron may have a thousand synapses, this makes our brains equivalent to a massively parallel computer comprised of 100 billion processors [1]. So, what does all this have to do with photoreceptors? The rods and cones are the cells of the visual system that create the first change in membrane potential, and they then communicate this information to the next cells via their synapse. The RPE regenerates the visual pigment, allowing the photoreceptors to continue functioning over and over. Blind Spot There is a unique spot in each eye where there are no photoreceptors of any type. This spot is where the optic nerve exits the eye and where the blood vessels that supply the retina enter and leave. The region is called the optic disk, or the blind spot, because we cannot see anything in that area. It is located nasal of the fovea by several millimeters and thus is in the opposite side of the visual field for each eye. The optic disk is centered about 4–5 mm from the fovea and is typically about 1.5–2 mm in diameter. It is not a perfect circle and varies somewhat in shape from person to person. It is necessary for our high-acuity vision as this is the port through which blood flows to nourish the metabolically active retina and information leaves from the eye. But, in that region we see no light. How is it that we do not notice this absence of vision [1, 2]? First, we have binocular vision, meaning that each eye is conveying information about the same visual scene, but from a slightly different vantage point. This, of course, allows us to determine the distance of various objects, but it also allows us to fill in the full information of a scene when a part of it is missing from one eye. Second, even when we are observing a scene with just one eye, the visual center of our brain fills in when there is information lacking. So, when we do not have information

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about a small space in our visual field, our brain fills this in with the same scene that we see in the adjacent areas. Thus, if it is a missing spot in the sky, the brain fills in the appropriate blue and continues the cloud pattern that is immediately adjacent. This means that if there is a single spot in an otherwise uniform field, and if that spot is only imaged onto the blind spot, it is invisible to us. You can test this directly with the diagram below. Close your left eye and focus on the cross with your right eye. Move this page closer and further from your eye until the mark on the right disappears. It should occur as you move this page slightly closer to your eye than reading distance.

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A great advantage of the optic disk and retinal arrangement is to give vertebrates higher acuity and speedier vision than would be otherwise possible. The reason is that the RPE, which provides essential biochemical regeneration necessary for photoreception, allows the photoreceptors to respond much more rapidly to changes in light than possible if they had to maintain their full biochemical function independently. And, the RPE provides that essential “black screen” behind our photoreceptors to retain our high visual acuity. Were the retina to be reversed, with photoreceptors in the inner retina, our acuity would likely be lessened and the speed of response reduced, and the profusion of blood vessels to provide nutrition for the added photoreceptor function would further limit this. Indeed, for invertebrates, for which the photoreceptors are first cells in the optical path, accommodations are made (such as having a compound eye, as seen in flies) that substantially restrict visual acuity. RETINAL PATHWAYS Through Pathway Once a rod or cone has transduced the light to a change in membrane potential, it then conveys this by its synapse onto a bipolar cell, with one group of bipolar cells receiving rod information and another receiving cone information. The bipolar cells are so named because they have processes going in two directions from their cell body. Their dendrites receive information from photoreceptors, and their axons transmit information to the ganglion cells. Bipolar cells thus span the inner half of the retina (see Fig. 1), from the outer plexiform layer (OPL; where the photoreceptors form their synaptic contacts onto bipolar and horizontal cells) to the inner plexiform layer (IPL; where the bipolar cells form synaptic contacts onto amacrine and ganglion cells). The cell bodies of the bipolar cells comprise most of the inner nuclear layer (INL), while the cell bodies of the photoreceptors comprise the outer nuclear layer (ONL) [4, 8]. Bipolar cells then synapse onto retinal ganglion cells in the IPL. The ganglion cell bodies comprise the ganglion cell layer (GCL), and their axons make the nerve fiber layer (NFL). This straight-through pathway also involves processing of the visual information. A significant part of this for the cone bipolar cells is to convert the signals into on and off signals, meaning responses to light onset or to light offset. So, half the bipolar cells

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are dedicated to transmitting an increase in signal when there is an increase in light in the region they subtend, while the other half of the cone bipolar cells convey a larger signal when there is a decrement in the light in the region they subtend. The first group is called ON bipolar cells, and the latter group is termed OFF bipolar cells simply to indicate they turn on or off with light, respectively. This on-and-off feature is conserved in transmission to the ganglion cells, so they also exhibit the same types of signals [4, 8, 9]. It is the ON pathway that facilitates our perception of light on a dark background; the OFF pathway provides the perception of dark images on a light background. These parallel channels to convey visual information are key in providing one other feature of our senses: the exquisite sensitivity to change. Like our other sensory systems, our visual system has the feature of observing any change in the visual image. One can readily observe this by staring at a constant image for tens of seconds. As you stare at one point in the image, the periphery gradually disappears and with time even the central portion of the image will fade. Anyone who has watched for meteors is especially aware of this; when you fixate on one star, all the others gradually disappear, but when a meteor (or airplane) crosses the sky it is vividly seen. Indeed, in reference to another point, when we then turn to look directly at the meteor, we see that it is much dimmer than when we saw it with our peripheral vision [4]. While cone bipolar cells are either ON or OFF, rod bipolar cells are all of the ON variety. Thus, for dim light, we are keenly aware when the light is on or increasing but less so when it is decreasing. Receptive Fields The visual field or receptive field of each photoreceptor is quite small. As discussed above, it is about 1 arc minute in size or the size of one pixel on an LCD monitor at normal viewing distance. In the central part of our vision, there is a correspondence of cones with their respective bipolar cells, so that this level of acuity is maintained. Thus, the receptive fields of cone bipolar cells approximate the receptive fields of the cone photoreceptors in the central fovea. But, as one moves further out, more and more cones synapse onto each bipolar cell, resulting in a lessening of acuity with distance from the fovea. Hence, the size of the receptive fields of bipolar cells is determined by the number of photoreceptors from which they receive input. Cone bipolar cell receptive fields are small, especially in the fovea, while rod bipolar cell receptive fields are much larger. One reason is that cones, in providing high-acuity vision, require each spot of light to be processed for us to maintain that resolution. The rod bipolars, which are used to determine if a dim light is present, combine the signals of many rods to increase the chance of seeing a very small light signal [1, 4]. Lateral Pathway Now, if we simply had the straight-through pathway from photoreceptors to bipolar cells to ganglion cells, we would have vision that is somewhat grainy, and we would not have the excellent discrimination of edges that is inherent in our vision. The two lateral paths in the retina are essential in rectifying this. The horizontal cells make lateral interactions in the outer retina, while the amacrine cells make lateral interactions in the inner retina.

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These laterally oriented cells are inhibitory, meaning that they provide negative feedback, or suppress the signals, to the adjacent cells. Thus, when a horizontal cell receives information of bright light from one photoreceptor, it emits a signal of darkness to the adjacent photoreceptors, lessening their light response. In any constant field of illumination, this simply decreases the overall response. But, in regions where there is a border of light and dark, this has the effect of increasing the lightness at the edge of the light region and increasing the darkness at the edge of the dark region. The result is contrast enhancement created by the outer retina, a feature that has been observed and re-created by many impressionist painters (one may readily call to mind the images of dancers by Edgar Dégas, in which the dark stage has a black line adjacent to a dancer, while the lightness of her arm or leg has a white line at its edge to denote the contrast enhancement perceived by the painter’s visual system). The other feature these laterally acting cells have is to create receptive fields in the bipolar and ganglion cells that are ON in the center and OFF in the surround for ON bipolar cells (and OFF center, ON surround for OFF bipolar cells). This is referred to as a center-surround receptive field, and it is the type of receptive field present for bipolar cells and ganglion cells. These center-surround receptive fields are due to the action of the horizontal cells, which connect to a large number of photoreceptors and thus make the larger inhibitory “surround” for each bipolar cell receptive field. Thus, the inhibitory feedback of the horizontal cells allows a sharpening of perception, calling to attention the contrasts inherent in our visual scenes [1, 4, 8]. Retinal Ganglion Cells The retinal ganglion cells, while being the third cell in the visual path (photoreceptor to bipolar to ganglion), also benefit from the lateral processing of the horizontal and amacrine cells. The horizontal feedback occurs in the outer retina, in the OPL, while the amacrine feedback occurs in the IPL. This IPL is a rich network of connections between bipolar, amacrine, and ganglion cells. And, it is spatially organized, with the OFF bipolar cells making synaptic contact in the outer half of the IPL and the ON bipolar cells making synaptic contact onto ganglion cell dendrites in the inner half of the IPL [4, 8, 9]. The ganglion cells that receive signals from ON bipolar cells are also ON center cells, having an OFF surround. The ones with OFF input are OFF center, ON surround ganglion cells. At this point in the retina, the visual information is much more processed, so the information that is transmitted to the vision center of the brain is not only of the color pixels we see, but also of the color contrast, contrast enhancement, and in some animals the information of the movement of objects. The ganglion cells are the innermost part of the retina; their axons join together at the optic disk and leave the eye by passing through the retina and sclera, creating the optic nerve. This nerve travels to the lateral geniculate nucleus of the thalamus, located in the center of the head. The ganglion cells are the first visual system cells to have a long axon, and they create action potentials to transmit their visual information from the eye. These action potentials thus carry information from the eye to the thalamus of the brain, where the information is then relayed to the visual cortex, which is located in the back of the brain in an area called the occipital lobe. It is in the visual cortex that we assemble the signals

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to find patterns and movements of those shapes as well as create the binocular image from the information sent by the two eyes [1]. Retinal Glia In addition to the neural cells of the retina, there are glial cells that span the thickness of the retina. These are primarily retinal Müller cells, and they serve several essential functions to maintain retinal function. The Müller cells extend from the photoreceptors to the ganglion cells and provide a biochemical support for retinal cells, removing excess neurotransmitter that has been released and restricting its diffusion. These cells also help balance the external ionic milieu around the neural cells, and they are implicated in other physiological processes. It is evident that when these cells or the RPE cells are not functioning properly in their physical and biochemical support, retinal neurons are subsequently damaged, and vision is attenuated or lost. This is the case for a variety of conditions and diseases of the eye [10]. REFERENCES 1. Dowling JE. The retina an approachable part of the brain. Harvard University Press, Cambridge, MA, 1987. 2. Walls GL. The vertebrate eye and its adaptive radiation. Hafner, New York, 1967. 3. Westheimer G. Specifying and controlling the optical image on the human retina. Prog Retin Eye Res 2006;25:19–42. 4. Kolb H. How the retina works. Am Sci 2003;91:28–35. 5. Fisher SK, Lewis GP, Linberg KA, Verado MR. Cellular remodeling in mammalian retina: results from studies of experimental retinal detachment. Prog Retin Eye Res 2005;24: 395–431. 6. Bharti K, Nguyen MT, Skuntz S, Bertuzzi S, Arnheiter H. The other pigment cell: specification and development of the pigmented epithelium of the vertebrate eye. Pigment Cell Res 2006;19:380–394. 7. Chen CK. The vertebrate phototransduction cascade: amplification and termination mechanisms. Rev Physiol Biochem Pharmacol 2005;154:101–121. 8. Lukasiewicz PD. Synaptic mechanisms that shape visual signaling at the inner retina. Prog Brain Res 2005;147:205–218. 9. Baccus SA. Timing and computation in inner retinal circuitry. Annu Rev Physiol 2007;69: 271–290. 10. Newman EA. Calcium increases in retinal glial cells evoked by light-induced neuronal activity. J Neurosci 2005;25:5502–5510.

2 Development of the Foveal Specialization Keely M. Bumsted O’Brien CONTENTS Introduction Foveal Development Conclusions and Perspectives References

INTRODUCTION “Vision is the foundation of intelligence and the chief source of our knowledge”[1]. Vision, our primary sensory modality, is supported by a complex anatomy and physiology that coordinates the interpretation of and interaction with our world. The initial steps in seeing begin in the retina, where the processing of important features such as color, form, and movement is initiated. These functions are mediated by interactions between a great diversity of cell types in the retina, the three glial and six major neuronal cell classes. The approximately 55 distinct retinal cell types are each nonrandomly distributed across the retina to maximize retinal coverage and are organized in characteristic topographic patterns. The most highly specialized area of our retina, which is also the focal point of cell topography, is located at the center of gaze in a region known as the macula leutea or macula. Within the macula is a morphologically distinct region called the fovea Centralis or fovea, which is responsible for the most acute color, spatial, and temporal visual resolution properties in primates [1–4]. The fovea is defined as a small anatomical pit or depression in the retina at the center of gaze; it lacks rod photoreceptors and has the highest density of cone photoreceptors. In humans, the fovea is responsible for our high visual acuity, measured to be 6/6 (20/20 or approximately 30 cycles per degree). Among mammals, the presence of a fovea is restricted to primates; however, many other vertebrates, such as many birds, certain species of fish, and chameleons, have a fovea and enhanced visual acuity [5–10]. All foveae share the common feature of a retinal pit, although many other characteristics, such as pit depth, width, cell composition, and density, are variable between species. In general, bird and fish foveae are smaller in diameter with steeper walls, leading to the description of the these foveae as convexiclivate [7, 11]. In addition to being steeper, many other vertebrate foveae contain a continuous inner nuclear layer (INL) and ganglion cell layer From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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(GCL) across the center of their pit, whereas these layers are missing in the human fovea [10]. In the pigeon (Columba livia), northern blue jay (Cyanocitta cristata), and ostrich (Struthio camelus) the fovea is shallower compared to the human, although visual acuity is close to, but slightly poorer than, human acuity under comparable conditions (approximately 16–20 cycles per degree) [10, 12–16]. Only the large birds of prey such as the wedge-tailed eagle (Aquila audax; 140 cycles per degree) have a higher visual acuity compared to primates. This increased acuity is likely based in the higher foveal cone density and has been suggested to be optically augmented by the steep structure of the pit [10, 13, 17]. The primate fovea is located in the center of the macula, an oval 1.5-mm wide yellow spot visible while viewing the fundus. The yellow color of the macula lutea is due to the presence of the carotenoid pigments lutein and zeaxanthin. Within the macular region lies the fovea (Fig. 1A, circle). The morphological features of the human fovea are characterized by a pit 600–800 µm wide that is lacking inner retinal neurons, blood vessels, and rod photoreceptors but contains the highest density of cones in the retina (Fig. 1B, bracketed region). Maximum foveal cone densities range from between about 100,000 and 400,000 cones/mm2 in humans (Fig. 1D; [18–24]). The very center of the fovea, the foveola, is further specialized in that it contains long- (L) and medium- (M) wavelength specific cone density peaks, while short- (S) wavelength specific cones, which elsewhere comprise 8–10% of the cone population, are absent from the central 20 µm of the foveola [3, 25]. Cone densities decline rapidly with eccentricity, such that even in the fovea there is a steep density gradient [20, 26]. Outside the region of high cone density, the edges of the fovea form a slope where the number of inner retinal neurons begins to increase as the foveal region gradually transitions to a fully layered retina at the rim margin (Fig. 1B, arrowheads). The retinal capillary network is present on the foveal rim but not on the foveal slope. The foveal rim is also distinguished by an accumulation of cells in the GCL and the INL [1, 2, 10, 26–28]. The accumulation of retinal ganglion cells on the foveal rim is correlated with the high density of cone photoreceptors in the foveal center (Fig. 1D) because each foveal cone stimulates two midget ganglion cells (i.e., one ON and one OFF). Each ON or OFF midget ganglion cell receives information from a single cone through an ON or OFF bipolar cell, respectively (Fig. 1C) [1, 29–31]. This dedicated line of information flow and the small size of the ganglion cell arbors and receptive fields in the central retina mean that individual foveal cones represent one small region of spatial information that is transmitted to the parvocellular layers of the dorsal lateral geniculate nucleus with little information loss (reviewed in [32]). Outside the fovea, there is a regular decrease in the density of cones, which is in parallel with the decrease in ganglion cell density and increased rod density (Fig. 1D). As the number of ganglion cells decreases, there is an increase in the size of ganglion cell dendritic arbors and receptive field sizes [33, 34]. Therefore, relative to the macular region, the peripheral retina has a greater summation of information due to the larger ganglion cell receptive field, lower cone densities, and increase in rod number [10, 20, 21, 35, 36]. The complex organization of the retina centered on the fovea is essential for our optimal visual functioning. When this topographical arrangement is compromised in the adult or during development, the result is poor vision. Almost a quarter of all adult

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Fig. 1. Characteristics of the adult human fovea. A Fundus photograph of a human macular region. The circled region indicates where the foveal pit is located. B Cross section of the human fovea stained with cresyl violet. In the center of the fovea (bracket), all the inner retinal neurons have been pushed aside, and the cones are found at a high density. The edges of the pit (arrowheads) contain a high density of ganglion cells and are where the edges of the avascular zone are located. C Diagram of a foveal cone, ON and OFF midget bipolar cell and ganglion cell circuit. Note the long axon of the foveal cone. D Photoreceptor densities across the human retina (modified from [18]). Cones peak in the center of the fovea and then fall quickly into the periphery. Rods are absent in the foveal center and peak outside the fovea. GC ganglion cell, GCL ganglion cell layer, INL inner nuclear layer, OD optic disk, ONL outer nuclear layer.

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visual impairment and many cases of childhood blindness are caused by defects in the fovea and “central” visual loss [37–47]. While the development of the human fovea has been previously reviewed, there has yet to be an overview specifically aimed at incorporating data from retinal development in nonprimate species with data available from primates. In this chapter, four key steps necessary for foveal development are defined and discussed: (1) the specification of foveal location at the center of gaze; (2) the generation of the rod-free zone; (3) the progressive increase in cone packing density and initial pit formation; and finally (4) the lengthy centrifugal displacement of the inner retinal cells (cells move away from the foveal center), forming the adult foveal pit and increase in cone density overlying the fovea. These steps must be executed in the correct sequence or foveal development will be stalled, resulting in foveal hypoplasia (discussed at the end of the chapter). FOVEAL DEVELOPMENT Foveal development in primates occurs over a protracted period, beginning before birth and then extending far into postnatal life. While the region of the retina in which the fovea will become established is the first to differentiate in early fetal life, establishment of adult-like characteristics of the fovea occurs a considerable time after birth in the final stages of retinal development. Therefore, the mechanisms that control the development of the foveal region must be tightly regulated over a long temporal sequence. The proper execution of each step is dependent on the success of the previous step. Any change in the normal progression of development will affect foveal structure and function (e.g., the degree of visual acuity). Specification of Foveal Location In the adult eye, the fovea is located 4.9 mm from the optic nerve head with little variability between eyes and individuals (Fig. 1A) [48]. These measures indicate that the placement of the fovea during development is tightly controlled and likely established early during retinal development. One possible scenario for placing the fovea would be that the foveal region is developing in a particular molecular environment that instructs progenitor cells to generate the unique properties of the macula. What are the signals that set up this environment? In the case of the fovea, first the location must be established. Using a chicken model, it has been shown that the specification of spatial location in the retina is determined as early as optic vesicle formation (in humans, this is approximately fetal week 3.4) [49–52]. Dividing cells located toward the distal tip of the optic vesicle has been shown by fate mapping lineage analysis techniques to give rise to central retinal regions, and cells in the anterior and posterior optic vesicle will give rise to the nasal and temporal retina, respectively [50, 52, 53]. These experiments did not take into account specific retinal location or photoreceptor topography. Further refinement of this regional map will allow the exact mapping of the future center of gaze. What controls the formation of the various axes and the central region of the retina? The address given to cells in different regions of the optic vesicle and optic cup is thought to be mediated by the expression of regionally specific genes, which segregate the developing eye into functionally distinct domains along the anterior posterior (AP) and dorsal ventral

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Fig. 2. Examples of compartmentalized gene expression in the developing eye. A The DV axis of the eye and early neural retina is subdivided into several domains based on the expression of specific genes. The dorsal retina is delineated by Tbx5, ephrin B2, ephrin B1, BMP 4, and RALDH. The ventral retina expresses Vax2, Pax2, RALDH 3, and RALDH6. B The temporal retina expresses Foxd1 (brain factor 2), CBF3, and Eph A3. Foxg1 (brain factor 1), SOHo-1, GH6, and CBF-1 delineate the nasal retinal region. The molecules that are specific for this region are CYP26, BMP2, and FGF8. D dorsal, N nasal, T temporal, V ventral. Modified from Schulte D, Bumsted-O’Brien KM. 2008.

(DV) axes [50, 52, 54–57]. Work from many groups has led to the creation of a regionally distinct topographic map (Fig. 2). The dorsal retina is delineated by a number of transcription factors and signaling molecules, including Tbx5, ephrin B2, ephrin B1, BMP 4, and RALDH1 [56–66], while the ventral retina expresses Vax2, Pax2, RALDH 3, and RALDH6 [55, 56, 59, 65, 67, 68] (Fig. 2A). The temporal retina is delineated Foxd1 (brain factor 2), CBF3, and Eph A3 [69, 70]. Foxg1 (brain factor 1), SOHo-1, GH6, and CBF-1 are segregated to the nasal retinal region [55–71] (Fig. 2B). There is some overlap of the nasal and temporal expression gradients; however, this does not seem to be important in setting up the photoreceptor patterning (Fig. 2B, white zone). All of the nasal/temporal restricted genes tested so far are involved in retinal ganglion cell pathfinding as manipulating expression does not perturb photoreceptor topography [56, 57, 72]. The dorsal and ventral expression patterns do not overlap along the horizontal meridian where the center of gaze is located; instead, there is a middle ground where neither dorsal nor ventral genes are expressed (Fig. 2A). The molecules that are specific for this region are CYP26, BMP2, BMP7, and FGF8 [73–75]. The expression of CYP26 allows for an abrupt step in the diffusion of Retinoic acid (RA) levels in the central retina. Overexpression of CYP26 induces a loss of ephrin B2 expression, a dorsal-associated gene [76]. It has been shown previously that the disturbance of ephrin B2 leads to a disorganization of the normal topographically organized retinal ganglion cell projections [57]. The expression of BMP2 along the horizontal meridian and BMP7 in the chicken area centralis may indicate that it could play a role in locating the region that will form the area centralis

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or fovea [73, 77]. Pax 6, an important regulatory gene in eye development, is expressed throughout the developing retina; however, the highest level of expression is in the central swath [78]. Overexpression of PAX6 in the developing eye extends the normally dorsally restricted domain of TBX5 and BMP4 into the ventral retina. PAX6 was shown to interact with VAX, leading to the modulation of PAX6 enhancer activity such that PAX6 function was inhibited in the ventral retina [78]. Collectively, these results lend strong support to the view that positional specification along both major retinal axes is already in place in the optic vesicle and early optic cup, a time before neurogenesis has been initiated. Thus, positional identities are assigned to the progenitor cells of the optic vesicle and optic cup long before the first postmitotic neurons differentiate in the retina. Formation of a Rod-Free Zone The next step in foveal development involves the formation of a rod-free zone. This region could be formed by either the generation of rods followed by selective cell death in the fovea or signaling (active or passive) to the progenitor cells to exclude the production of rods in the developing fovea. In this section, I argue that the latter mechanism is likely to be used in the formation of the human rod-free zone. Experiments in the chicken have provided evidence to support this argument. While chickens do not have a fovea, their retina contains a rod-free area centralis at the center of gaze that allows them to have an acuity of 7 cycles per degree [79, 80]. The chicken experiments also showed the importance of the regionally specific genes to the generation of the retinal axes and the specification of the rod-free zone. When Schulte and colleagues manipulated the ventrally expressed transcription factor VAX2, the rod-free zone was lost [56, 72]. Overexpression of the dorsal gene TBX5 causes local disturbances in the rod pattern but does not remove the rod-free zone [72]. This indicates that the boundaries set up by regional gene expression patterns are critical to localizing and specifying the rod-free zone of the central retinal region. The data in the chicken suggest that rods are actively excluded from the developing fovea. In humans, by the time that the optic vesicle invaginates to form the optic cup (fetal week 4.5), the developing retina, which lies on the inner side of the cup, has a dramatic expansion in the number of dividing cells [81]. Cell birth starts at the site of the future fovea and then spreads from the retinal center toward the periphery in a wave of cell birth. There is no clear evidence regarding when ganglion cells are generated in the human. In a morphological study, Mann reported the appearance of the first axons in the optic nerve at fetal week 7 [28]. An estimation of human cell birth can be extrapolated from cell birth dating data obtained in the developing monkey. These data suggest that the first cells to exit the cell cycle in humans, the retinal ganglion cells, appear around fetal week 7 in the foveal region [81–84]. When correlated with morphology, the data indicate that retinal ganglion cells are born slightly before fetal week 7. The remaining retinal cell types are born in an orderly sequence beginning in the fovea and then spreading out into the periphery. Horizontal cells are born shortly after ganglion cells in the incipient fovea between fetal weeks 7 and 8, followed by cone photoreceptors. Next, amacrine cells and bipolar cells become postmitotic. The last cell types to be generated are the rod photoreceptors and Müller glia [84]. All cells are postmitotic in the fovea by fetal week 10. In the periphery, the last cells have been generated by fetal week 30 [85].

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Fig. 3. Development of the rod-free zone. A Fetal week 11 human fovea with short cuboidal cones in the outer nuclear layer. Rods are not present in the fovea. The inner nuclear layer (INL) and ganglion cell layer (GCL) with the intervening synaptic layers are present in the incipient fovea. B Fetal week 11 edge of the fovea. Rods begin to be detected (arrows) on the edge of the fovea. C Rods on the edge of the fovea are labeled with an antibody to NR2E3 (arrows) and D Nrl (arrows) (modified from [89]). D The location of the rod-free zone (pale shading) in relation to the rod-dominant retina (dark shading). E The unknown signals (?) working on the progenitor cell (PC) to produce rods and a rod-free zone are indicated in the diagram to the right of the retina diagram. OD optic disk.

From the earliest point of identification, fetal week 11, rod photoreceptors are missing from the foveal center (Fig. 3A). Rods are first observed on the foveal edge (Fig. 3B–D, arrows). This lack of rods in the fovea in the human appears to be intricately linked to cell generation rather than resulting from the death of inappropriately generated rods. During the development of the fovea, it has been shown that there is little cell death in the human photoreceptor layer, indicating that it is unlikely that excess photoreceptors are generated then eliminated by cell death [86, 87]. In addition, molecules associated with rod differentiation, Nrl and NR2E3, are never detected in the foveal region, suggesting that rods are developmentally excluded from the fovea [88, 89] (Fig. 3C,D).

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Therefore, in the case of rods it seems that they are never generated in the fovea center. While there has been progress investigating the inductive influences on progenitor cells to stimulate rod differentiation [89, 90], the signals that control the exclusive differentiation of cones remain to be established. Therefore, the first two steps of foveal development are intricately linked, although the exact mechanism utilized to establish foveal location and exclude rods remains elusive (Fig. 3E). Cones, Ganglion Cells, and Initial Pit Formation The lack of rods in the fovea appears to be an early developmental patterning event; however, other characteristics of the adult fovea such as the accumulation of a high cone photoreceptor density and the formation of the pit appear to be later developmental events. When the foveal region becomes identifiable at fetal week 11, foveal neurons are postmitotic, synapses have been formed, all the retinal layers are present, and foveal cones can be identified as short, fat cuboidal cells lying in a single layer at a density of 11,200 cones/mm2 (Fig. 4A) [2, 26, 83, 85, 91, 92]. Which forces act on the foveal cones to influence them to change from a cuboidal shape to the thin, elongated cone in the mature fovea? The first observable change in the foveal cone is the elongation of the axonal process (Figs. 4 and 5). This alteration of cell shape is necessary for the cones to maintain their synaptic connections as they pack tightly into the fovea to reach the average adult cone density of approximately 200,000 cones/mm2 [3]. The exact mechanism by which the cones begin to change their morphology such that they can pack more tightly in the center of the fovea is unknown; however, it has been proposed that the mechanism for this elongation is the differential expression of the fibroblast growth factor (FGF) family members and their receptors on foveal cones [93, 94]. FGF signaling may mediate the morphological cell shape changes of foveal cones that are associated with increased density in the foveal cone mosaic in the early phase of photoreceptor accumulation (Fig. 5) [92, 95]. In humans at Fetal week 14, the maximum spatial density of foveal ganglion cells is approximately 22,500 cells/mm2, and this density increases to about 31,500 cells/mm2 at Fetal weeks 16–17 [85]. Therefore, at the same time that the cones are changing shape, the ganglion cells begin to accumulate in the incipient foveal region (Fig. 4B, arrow) [28]. The timing of this peak in maximum ganglion cell density corresponds with the appearance of a distinctive dome in the GCL (Fig. 4B,C, arrow) [27, 28, 91]. Once the GCL reaches a critical thickness (approximately 7–9 cells thick), the initial formation of the pit is initiated between fetal weeks 24 and 28 (Fig. 4C,D, arrow) [28]. The beginning of the foveal pit begins as a shallow depression and is observed as a progressive decrease in the depth of the inner retinal layers [2]. By fetal week 28, the fovea contains a clearly defined pit with ganglion cells and inner retinal neurons beginning to move away from the foveal center (Fig. 4D, arrow). The foveal cones are further elongating, their pedicles are being pulled away from the foveal center, and the density of foveal cones increases to 22,268 cells/mm2 [26, 92]. The cells of the inner and outer nuclear layers move away from the foveal center (centrifugal migration), while at the same time foveal cones translocate in the opposite direction (centripetal), becoming more densely packed (18,500 cones/mm2) (Fig. 4E) [26, 92]. It is at this point that the third stage of foveal development is completed. In the next stage, there are additional influences on the

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Fig. 4. Morphological development of the human fovea. A A cross section of the fovea at fetal week 11 stained with cresyl violet. There are no rods present in the foveal center. B Cross section of a fetal week 16 human fovea stained with cresyl violet. Note the cuboidal shape of the foveal cones. C Fetal week 24 foveal retinal cross section. The arrow indicates the center of the developing fovea and where there is an accumulation of ganglion cells in the foveal center in the region where the pit will form. D A detailed drawing of a fetal week 28 foveal cross section. This is the first age at which an indentation in the fovea is observed (arrow). E Drawing of a fetal week 32 foveal cross section. The pit is becoming deeper and the inner retinal layers thinning. Note the length of the foveal cones compared to those in A. F Postnatal 8 week drawing of a foveal retinal cross section. The foveal pit is much deeper compared to fetal week 32, and the foveal cone axons have significantly elongated in the foveal center (arrow). (A and B modified from [91].) (C, D, E, and F modified from high-resolution scans from [27].) The arrows indicate the center of the developing fovea.

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Fig. 5. Drawing of the central foveal cones throughout development. At fetal week 11, the cones are cuboidal and then begin to slowly elongate from fetal weeks 16, 20, and 24. These ages correspond with the increase in cone packing that may be mediated by fibroblast growth factor (FGF) signaling. By fetal week 32, the cones begin to elongate further, such that after birth, they are significantly thinner with longer axons compared with fetal week 11. These cones are changing shape due to proposed mechanical influences that form the deep foveal pit. (Cones were traced from images in Fig. 4 and from images from [91].)

forming foveal region that mediate the further increase in cone cell density and complete pit formation. Retinal blood vessels are first detected emerging from the optic disk at approximately fetal week 14 and rapidly spread into the superior and inferior quadrants of the retina; however, the vascularization of the macular region is delayed. By fetal week 22, inner retinal vessels have grown out to cover greater than 50% of the retina, but the ring of vessels that will form the avascular zone in the adult has not yet begun to encircle the central retina. This ring of vasculature starts forming between fetal weeks 25 and 28, but the precise timing of the formation of the foveal avascular zone in humans has not been established. Interestingly, the foveal region is never vascularized during normal development, suggesting that there is a repulsive cue acting to repel the vessels [96–98]. There does seem to be a correlation between the formation of the avascular zone and the formation of the fovea; however, this has yet to be tested. A role for the vasculature in the formation of mature foveal pit is discussed in the next section. Deep Foveal Pit Formation The next stage of foveal development is the refinement of the pit to exclude all nonphotoreceptor cells and increase cone photoreceptor numbers. The initial increase in cone density follows a shallow slope; however, between fetal week 32 and birth, the slope flattens (Figs. 4E and 6). After birth, there is a marked change in the slope such that the accumulation of cones is accelerated (Fig. 4F). As cells of the inner and outer nuclear layers are displaced, they maintain their initial synaptic relationships, which are critical for the maintenance of acuity in the one–one circuitry of the midget system (Figs. 1C, 4F, and 5).

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The mechanism of deep foveal pit formation is not yet known. It is possible that these changes are related to the completion of the ring of vasculature around the fovea. The pit becomes progressively deeper and more pronounced through the rest of fetal development and early childhood. The length of the cone axons continues to increase, and the cell bodies become thinner (Fig. 5). The final adult cone density peaks at more than 200,000 cones/mm2 [2, 26, 28, 99]. There has been considerable debate concerning the mechanism of pit formation. This is because it is difficult to test the hypotheses since an animal model has not been available that has a foveal pit and is easily manipulated during retinal development. A recent model for foveal pit formation was proposed by Springer and Hendrickson and based on morphological observations in the developing human retina [100]. They applied a virtual engineering model combined with finite element analysis to identify mechanical mechanisms important for pit formation. They hypothesized that the pit emerges within the foveal region because it contains an avascular zone, which makes this region more elastic compared to surrounding vascularized retina (Fig. 4 E,F, arrowheads). The conclusion from their modeling study is that once differential elasticity is established by the avascular zone, pit formation and a concomitant increase in cone density can be driven by either intraocular pressure or ocular growth-induced retinal stretch. Foveal Hypoplasia The long developmental time frame of foveal development allows for numerous points where mutations or missteps can occur. These mistakes or defects can lead to foveal hypoplasia, a condition in which an individual lacks a foveal pit, avascular zone, and rod-free zone [41]. Foveal hypoplasia results in decreased vision with reduced acuity (6/24 to 6/120 or 20/80 to 20/400 [41, 101]). The underlying cause of foveal hypoplasia is not yet known, although it has been associated with many eye conditions, including albinism, microcornea, familial and presenile cataracts, and PAX6 mutations [38, 39, 44]. The degree of hypoplasia is likely correlated with the gene defect. For example, it has been suggested that PAX6 is associated with the specification of foveal location. If foveal location is not established (step 1), then the correct microenvironment for setting up the rod-free zone and pit formation will be missing, thus halting development at an early time point. In the case of albinism, the central retina is lacking a rod-free zone, which suggests that this step is critical to stimulate a fovea [41]. It may be that the presence of vasculature across the foveal region also inhibits pit formation. Taken together, the characteristics of foveal hypoplasia suggest that interruption of any of the steps outlined in this chapter will cause moderate-to-severe central retinal malformations that have a profound effect on functional vision (Fig. 6). CONCLUSIONS AND PERSPECTIVES Changes in the foveal cone photoreceptors and the progression of primate foveal pit formation are summarized in Figs. 5 and 6. These figures show the progression of the different steps of foveal development in relation to major cellular movements and developmental events. Step 1 is initiated very early in retinal development (approximately fetal week 3). Once the location is specified, the correct cellular environment must be

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Fig. 6. Summary of the stages of foveal development. The steps in foveal development are outlined with reference to the change in cone density, developmental age, and important morphological events. Fwk fetal week OD optic disk (Modified from [104].)

specified during the second step, which is completed by fetal week 11. Step 3 slowly increases the density of cones via signaling molecules such as the FGFs (Figs. 5 and 6). Step 3 is completed around fetal week 32. Once the avascular ring is completed around birth, there is a sharp rise in the density of cones and a concomitant increase in cone length and decrease in cone diameter (Figs. 5 and 6). The mature fovea is apparent at approximately 4 years of age [2, 26]. When the structure of the human foveal pit is compared to the deep foveal pits of fish, birds, and lizards, it is difficult to imagine that they are formed by a similar mechanism. For example, one of the requirements of the virtual engineering model is the presence of an avascular zone. As none of these animal models has retinal vasculature, any forces proposed to be acting on the avacular zone to form a deep pit would be missing. Therefore, step 4 may not be required for the formation of all foveae, but this restriction does not have to be placed on the other preceding three steps when investigating a mechanism for early foveal development in nonprimates. There may be a universal mechanism in place to carry out the initial steps in foveal formation, such as in step 1, the specification of location. All of the observed foveal pits are faithfully located in a particular retinal location, indicating that the placement of the fovea at the center of gaze is an important step in the formation of the fovea. The development of the rod-free zone (step 2) also appears to be a generally conserved foveal requirement. The majority of foveae studied have a clearly defined rod-free zone in the fovea [6–8, 102, 103]. An increased cone density (step 3) appears to be a common feature in the center of foveal pits. This suggests that it is possible that the mechanisms that function early in foveal development are conserved among species to create a rod-free, cone-rich foveal pit, but that the formation of either a deep, steep foveal pit or a wide, shallow pit (primates) is dependent on divergent mechanisms. The argument concerning the evolution of the fovea can be summarized in the question: Is there can be an evolutionarily conserved mechanism to create the foveal pit in all these creatures, or does the distant relation of the primates, birds, and lizards

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require that a completely separate mechanism exist? Likely, the answer lies somewhere in the middle. The primate fovea can be characterized by the modification of retinal lamination, specialization of photoreceptors, and variation of the retinal vascular pattern. All of these characteristics combine to mediate our high visual acuity. The development of the fovea begins early in fetal development and continues until at least 4 years of age. When the four steps outlined in this chapter are faithfully reproduced, a characteristic fovea is formed (Figs. 1B and 5). Future research will help to elucidate the mechanisms by which the foveal specialization is formed with the hope of recapitulating these events in congenitally malformed or diseased retinae. ACKNOWLEDGMENTS I would like to thank Drs. Jan Provis, Dorothea Schulte, and Anita Hendrickson for valuable discussions of this topic. A special thanks is extended to Dr. Provis for her critical reading of the manuscript. Research in my labs is supported by the Center for Vision Excellence, the University of Auckland Staff Research Fund, and the Auckland Medical Research Foundation. REFERENCES 1. Polyak S. The retina. Chicago: University of Chicago Press; 1941. 2. Hendrickson AE, Yuodelis C. The morphological development of the human fovea. Ophthalmology 1984;91;603–612. 3. Curcio CA, et al. Distribution and morphology of human cone photoreceptors stained with anti-blue opsin. J Comp Neurol 1991;312;610–624. 4. Robinson, S. Development of the mammalian retina. In: Dreher B, Robinson S, eds. Neuroanatomy of the visual pathways and their development. Boca Raton, FL: CRC Press; 1991:69–128. 5. Collin SP, Collin HB. Topographic analysis of the retinal ganglion cell layer and optic nerve in the sandlance Limnichthyes fasciatus (Creeiidae, Perciformes). J Comp Neurol 1988;278:226–241. 6. Collin SP, Collin HB. The morphology of the retina and lens of the sandlance, Limnichthyes fasciatus (Creeiidae). Exp Biol 1988;47:209–218. 7. Collin SP, Collin HB. The foveal photoreceptor mosaic in the pipefish, Corythoichthyes paxtoni (Syngnathidae, Teleostei). Histol Histopathol 1999;14:369–382. 8. Collin SP, Lloyd DJ, Wagner HJ. Foveate vision in deep-sea teleosts: a comparison of primary visual and olfactory inputs. Philos Trans R Soc Lond B Biol Sci 2000;355:1315–1320. 9. Fritsches KA, Marshall NJ. Independent and conjugate eye movements during optokinesis in teleost fish. J Exp Biol 2002;205:1241–1252. 10. Walls GL. The vertebrate eye and its adaptive radiation. Bloomfield Hills, MI: Cranbrook Institute of Science; 1942. 11. Kirk EC, Kay RF. The evolution of high visual acuity in the Anthropoidea. In: Ross CF, Kay RF, eds. Anthropoid origins: new visions, New York: Kluwer /Plenum; 2004:539–602. 12. Boire D, Dufour JS, Theoret H, Ptito M. Quantitative analysis of the retinal ganglion cell layer in the ostrich, Struthio camelus. Brain Behav Evol 2001;58:343–355. 13. Fite KV, Rosenfield-Wessels S. A comparative study of deep avian foveas. Brain Behav Evol 1975;12:97–115.

30

Bumsted O’Brien

14. Hodos W, Miller RF, Fite KV. Age-dependent changes in visual acuity and retinal morphology in pigeons. Vision Res 1991;31:669–677. 15. Blough PM. The visual acuity of the pigeon for distant targets. J Exp Anal Behav 1971;15:57–67. 16. Hodos W, Leibowitz RW, Bonbright JC Jr. Near-field visual acuity of pigeons: effects of head location and stimulus luminance. J Exp Anal Behav 1976;25:129–141. 17. Reymond L. Spatial visual acuity of the eagle Aquila audax: a behavioural, optical and anatomical investigation. Vision Res 1985;25:1477–1491. 18. Osterberg G. Topography of the layer of rods and cones in the human retina. Acta Ophthal 1935;6:1–103. 19. Ahnelt PK, Kolb H, Pflug R. Identification of a subtype of cone photoreceptor, likely to be blue sensitive, in the human retina. J Comp Neurol 1987;255:18–34. 20. Curcio CA, Sloan KR, Kalina RE, Hendrickson AE. Human photoreceptor topography. J Comp Neurol 1990;292:497–523. 21. Wikler KC, Williams RW, Rakic P. Photoreceptor mosaic: number and distribution of rods and cones in the rhesus monkey retina. J Comp Neurol 1990;297:499–508. 22. Diaz-Araya CM, Provis JM, Billson FA. NADPH-diaphorase histochemistry reveals cone distributions in adult human retinae. Aust N Z J Ophthalmol 1993;21:171–179. 23. Perry VH, Cowey A. The ganglion cell and cone distributions in the monkey’s retina: implications for central magnification factors. Vision Res 1985;25:1795–1810. 24. Packer O, Hendrickson AE, Curcio CA. Photoreceptor topography of the retina in the adult pigtail macaque (Macaca nemestrina). J Comp Neurol 1989;288:165–183. 25. Bumsted K, Hendrickson A. Distribution and development of short-wavelength cones differ between Macaca monkey and human fovea. J Comp Neurol 1999;403:502–516. 26. Yuodelis C, Hendrickson A. A qualitative and quantitative analysis of the human fovea during development. Vision Res 1986;26:847–855. 27. Bach L, Seefelder R. Entwicklungsgeschichte des Menchlichlichen Auges. Leipzig, 1911, 1912, 1914. 28. Mann, I. In: The development of the human eye. 2nd ed. New York: Grune and Stratton; 1964. 29. Dowling JE, Boycott BB. Retinal ganglion cells: a correlation of anatomical and physiological approaches. UCLA Forum Med Sci 1969;8:145–161. 30. Calkins DJ, Tsukamoto Y, Sterling P. Microcircuitry and mosaic of a blue-yellow ganglion cell in the primate retina. J Neurosci 1998;18:3373–3385. 31. Calkins DJ, Schein SJ, Tsukamoto Y, Sterling P. M and L cones in macaque fovea connect to midget ganglion cells by different numbers of excitatory synapses. Nature 1994;371:70–72. 32. Wassle H. Parallel processing in the mammalian retina. Nat Rev Neurosci 2004;5:747–757. 33. Ghosh KK, Goodchild AK, Sefton AE, Martin PR. Morphology of retinal ganglion cells in a new world monkey, the marmoset Callithrix jacchus. J Comp Neurol 1996;366:76–92. 34. Goodchild AK, Ghosh KK, Martin PR. Comparison of photoreceptor spatial density and ganglion cell morphology in the retina of human, macaque monkey, cat, and the marmoset Callithrix jacchus. J Comp Neurol 1996;366:55–75. 35. DeBruyn EJ, Wise VL, Casagrande VA. The size and topographic arrangement of retinal ganglion cells in the galago. Vision Res 1980;20:315–27. 36. Rohen JW, Castenholz A. [On the centralization of the retina in primates]. Folia Primatol (Basel) 1967;5:92–147. 37. Constable IJ. Age-related macular degeneration and its possible prevention. Despite well publicised claims of the therapeutic value of dietary supplements and other new treatments, the evidence for their effectiveness is modest. Med J Aust 2004;181:471–472. 38. Curran RE, Robb RM. Isolated foveal hypoplasia. Arch Ophthalmol 1976;94:48–50.

Development of the Foveal Specialization

31

39. Oliver MD, Dotan SA, Chemke J, Abraham FA. Isolated foveal hypoplasia. Br J Ophthalmol 1987;71:926–930. 40. Haefemeyer JW, Knuth JL. Albinism. J Ophthalmic Nurs Technol 1991;10:55–62. 41. Mietz H, Green WR, Wolff SM, Abundo GP. Foveal hypoplasia in complete oculocutaneous albinism. A histopathologic study. Retina 1992;12:254–260. 42. Azuma N, Nishina S, Yanagisawa H, Okuyama T, Yamada M. PAX6 missense mutation in isolated foveal hypoplasia. Nat Genet 1996;13:141–142. 43. Meyer CH, Lapolice DJ, Freedman SF. Foveal hypoplasia in oculocutaneous albinism demonstrated by optical coherence tomography. Am J Ophthalmol 2002;133:409–410. 44. McGuire DE, Weinreb RN, Goldbaum MH. Foveal hypoplasia demonstrated in vivo with optical coherence tomography. Am J Ophthalmol 2003;135:112–114. 45. Vincent MC, et al. Variable phenotype related to a novel PAX 6 mutation (IVS4+5G>C) in a family presenting congenital nystagmus and foveal hypoplasia. Am J Ophthalmol 2004;138:1016–1021. 46. Pal B, et al. A new phenotype of recessively inherited foveal hypoplasia and anterior segment dysgenesis maps to a locus on chromosome 16q23.2–24.2. J Med Genet 2004;41:772–777. 47. Azuma N, et al. The Pax6 isoform bearing an alternative spliced exon promotes the development of the neural retinal structure. Hum Mol Genet 2005;14:735–745. 48. Williams TD, Wilkinson JM. Position of the fovea centralis with respect to the optic nerve head. Optom Vis Sci 1992;69:369–377. 49. Dutting D, Meyer SU. Transplantations of the chick eye anlage reveal an early determination of nasotemporal polarity. Int J Dev Biol 1995;39:921–931. 50. Dutting D, Thanos S. Early determination of nasal-temporal retinotopic specificity in the eye anlage of the chick embryo. Dev Biol 1995;167:263–281. 51. Thanos S, Dutting D. Plasticity in the developing chick visual system: topography and maintenance of experimentally induced ipsilateral projections. J Comp Neurol 1988;278:303–311. 52. Thanos S, Mey J, Dutting D, Hummler E. Positional determination of the naso-temporal retinal axis coincides with asymmetric expression of proteins along the anterior-posterior axis of the eye primordium. Exp Eye Res 1996;63:479–492. 53. Mueller BK, Dutting D, Haase A, Feucht A, Macchi P. Partial respecification of nasotemporal polarity in double-temporal chick and chimeric chick-quail eyes. Mech Dev 1998;74:15–28. 54. Peters MA. Patterning the neural retina. Curr Opin Neurobiol 2002;12:43–48. 55. Peters MA, Cepko CL. The dorsal-ventral axis of the neural retina is divided into multiple domains of restricted gene expression which exhibit features of lineage compartments. Dev Biol 2002;251:59–73. 56. Schulte D, Furukawa T, Peters MA, Kozak CA, Cepko CL. Misexpression of the Emx-related homeobox genes cVax and mVax2 ventralizes the retina and perturbs the retinotectal map. Neuron 1999;24:541–553. 57. Schulte D, Cepko CL. Two homeobox genes define the domain of EphA3 expression in the developing chick retina. Development 2000;127:5033–5045. 58. Logan M, Simon HG, Tabin C. Differential regulation of T-box and homeobox transcription factors suggests roles in controlling chick limb-type identity. Development 1998;125:2825–2835. 59. Uemonsa T, Sakagami K, Yasuda K, Araki M. Development of dorsal-ventral polarity in the optic vesicle and its presumptive role in eye morphogenesis as shown by embryonic transplantation and in ovo explant culturing. Dev Biol 2002;248:319. 60. Dutting D, Handwerker C, Drescher U. Topographic targeting and pathfinding errors of retinal axons following overexpression of ephrinA ligands on retinal ganglion cell axons. Dev Biol 1999;216:297–311. 61. Feldheim DA, et al. Loss-of-function analysis of EphA receptors in retinotectal mapping. J Neurosci 2004;24:2542–2550.

32

Bumsted O’Brien

62. Zhang XM, Yang XJ. Temporal and spatial effects of Sonic hedgehog signaling in chick eye morphogenesis. Dev Biol 2001;233:271–290. 63. Li H, et al. A retinoic acid synthesizing enzyme in ventral retina and telencephalon of the embryonic mouse. Mech Dev 2000;95:283–289. 64. Niederreither K, Vermot J, Schuhbaur B, Chambon P, Dolle P. Retinoic acid synthesis and hindbrain patterning in the mouse embryo. Development 2000;127:75–85. 65. Suzuki R, et al. Identification of RALDH-3, a novel retinaldehyde dehydrogenase, expressed in the ventral region of the retina. Mech Dev 2000;98:37–50. 66. Zhao D, et al. Molecular identification of a major retinoic-acid-synthesizing enzyme, a retinaldehyde-specific dehydrogenase. Eur J Biochem 1996;240:15–22. 67. Barbieri AM, et al. A homeobox gene, vax2, controls the patterning of the eye dorsoventral axis. Proc Natl Acad Sci U S A 1999;96:10729–10734. 68. Ohsaki K, Morimitsu T, Ishida Y, Kominami R, Takahashi N. Expression of the Vax family homeobox genes suggests multiple roles in eye development. Genes Cells 1999;4:267–276. 69. Shintani T, et al. Large-scale identification and characterization of genes with asymmetric expression patterns in the developing chick retina. J Neurobiol 2004;59:34–47. 70. Yuasa J, Hirano S, Yamagata M, Noda M. Visual projection map specified by topographic expression of transcription factors in the retina. Nature 1996;382:632–635. 71. Deitcher DL, Fekete DM, Cepko CL. Asymmetric expression of a novel homeobox gene in vertebrate sensory organs. J Neurosci 1994;14:486–498. 72. Schulte D, Peters MA, Sen J, Cepko CL. The rod photoreceptor pattern is set at the optic vesicle stage and requires spatially restricted cVax expression. J Neurosci 2005;25:2823–2831. 73. Belecky-Adams T, Adler R. Developmental expression patterns of bone morphogenetic proteins, receptors, and binding proteins in the chick retina. J Comp Neurol 2001;430:562–572. 74. Martinez-Morales JR, et al. Differentiation of the vertebrate retina is coordinated by an FGF signaling center. Dev Cell 2005;8:565–574. 75. Sakai Y, Luo T, McCaffery P, Hamada H, Drager UC. CYP26A1 and CYP26C1 cooperate in degrading retinoic acid within the equatorial retina during later eye development. Dev Biol 2004;276:143–157. 76. Sen J, Harpavat S, Peters MA, Cepko CL. Retinoic acid regulates the expression of dorsoventral topographic guidance molecules in the chick retina. Development 2005;132:5147–5159. 77. Adler R, Belecky-Adams TL. The role of bone morphogenetic proteins in the differentiation of the ventral optic cup. Development 2002;129:3161–3171. 78. Leconte L, Lecoin L, Martin P, Saule S. Pax6 interacts with cVax and Tbx5 to establish the dorsoventral boundary of the developing eye. J Biol Chem 2004;279:47272–47277. 79. Schmid KL, Wildsoet CF. Assessment of visual acuity and contrast sensitivity in the chick using an optokinetic nystagmus paradigm. Vision Res 1998;38:2629–2634. 80. Morris VB. An afoveate area centralis in the chick retina. J Comp Neurol 1982;210:198–203. 81. Barishak Y. Embryology of the eye and its adnexae. New York: Karger; 1992. 82. Hendrickson AE. Synaptic development in macaque monkey retina and its implications for other developmental sequences. Perspect Dev Neurobiol 1996;3:195–201. 83. Hendrickson AE. Primate foveal development: a microcosm of current questions in neurobiology. Invest Ophthalmol Vis Sci 1994;35:3129–3133. 84. Rapaport DH, Fletcher JT, LaVail MM, Rakic P. Genesis of neurons in the retinal ganglion cell layer of the monkey. J Comp Neurol 1992;322:577–588. 85. Provis JM, van Driel D, Billson FA, Russell P. Development of the human retina: patterns of cell distribution and redistribution in the ganglion cell layer. J Comp Neurol 1985;233:429–451.

Development of the Foveal Specialization

33

86. Georges P, Madigan MC, Provis JM. Apoptosis during development of the human retina: relationship to foveal development and retinal synaptogenesis. J Comp Neurol 1999;413:198–208. 87. Penfold PL, Provis JM. Cell death in the development of the human retina: phagocytosis of pyknotic and apoptotic bodies by retinal cells. Graefes Arch Clin Exp Ophthalmol 1986;224:549–553. 88. Bumsted O’Brien K, Schulte D, Hendrickson A. Expression of photoreceptor-associated molecules during fetal human eye development. Mol Vis 2003;9:401–409. 89. Bumsted O’Brien KM, et al. Expression of photoreceptor-specific nuclear receptor NR2E3 in rod photoreceptors of fetal human retina. Invest Ophthalmol Vis Sci 2004;45:2807–2812. 90. Mears AJ, et al. Nrl is required for rod photoreceptor development. Nat Genet 2001;29: 447–452. 91. Hendrickson A, Provis JM. Comparison of development of the primate fovea centralis with peripheral retina. In: Sernagor E, Eglen S, Harris WA, Wong R, eds. Retinal development. Cambridge, UK: Cambridge University Press; 2006:126–149. 92. Diaz-Araya C, Provis JM. Evidence of photoreceptor migration during early foveal development: a quantitative analysis of human fetal retinae. Vis Neurosci 1992;8:505–514. 93. Cornish EE, et al. Gradients of cone differentiation and FGF expression during development of the foveal depression in macaque retina. Vis Neurosci 2005;22:447–459. 94. Cornish EE, Natoli RC, Hendrickson A, Provis JM. Differential distribution of fibroblast growth factor receptors (FGFRs) on foveal cones: FGFR-4 is an early marker of cone photoreceptors. Mol Vis 2004;10:1–14. 95. Springer AD, Hendrickson AE. Development of the primate area of high acuity, 3: temporal relationships between pit formation, retinal elongation and cone packing. Vis Neurosci 2005;22:171–185. 96. Provis JM, et al. Development of the human retinal vasculature: cellular relations and VEGF expression. Exp Eye Res 1997;65:555–568. 97. Gariano RF, Sage EH, Kaplan HJ, Hendrickson AE. Development of astrocytes and their relation to blood vessels in fetal monkey retina. Invest Ophthalmol Vis Sci 1996;37:2367–2375. 98. Gariano RF, Iruela-Arispe ML, Sage EH, Hendrickson AE. Immunohistochemical characterization of developing and mature primate retinal blood vessels. Invest Ophthalmol Vis Sci 1996;37:93–103. 99. Provis JM, Billson FA, Russell P. Ganglion cell topography in human fetal retinae. Invest Ophthalmol Vis Sci 1983;24:1316–1320. 100. Springer AD, Hendrickson AE. Development of the primate area of high acuity. 1. Use of finite element analysis models to identify mechanical variables affecting pit formation. Vis Neurosci 2004;21:53–62. 101. Health MO. Children with disabilities. In: English B, ed. Our children’s health: key findings on the health of New Zealand children. Wellington, NZ: Ministry of Health; 1998:51–65. 102. Collin SP. Specialisations of the teleost visual system: adaptive diversity from shallowwater to deep-sea. Acta Physiol Scand Suppl 1997;638:5–24. 103. Collin SP. Topography and morphology of retinal ganglion cells in the coral trout Plectropoma leopardus (Serranidae): a retrograde cobaltous-lysine study. J Comp Neurol 1989;281:143–158. 104. Provis JM, Penfold PL, Cornish EE, Sandercoe TM, Madigan MC. Anatomy and development of the macula: specialisation and the vulnerability to macular degeneration. Clin Exp Optom 2005;88:269–281.

3 An Update on the Regulation of Rod Photoreceptor Development Edward M. Levine and Sabine Fuhrmann CONTENTS Introduction Brief Overview of Retinal Development and Early Stages of Rod Photoreceptor Differentiation Transcription Factors Extracellular Factors and Signal Transduction Pathways Conclusions and Future Prospects References

INTRODUCTION Photoreceptors are the most abundant retinal cells in all vertebrate species, and rods significantly outnumber cones in humans. Many retinal degenerative diseases arise due to perturbations in rod photoreceptor physiology or survival. Replacing dying photoreceptors by de novo production of new photoreceptors from a stem cell source is a strategy much sought after for treating these diseases. This approach in its essence requires our ability to manipulate stem cells and recapitulate the process of rod photoreceptor development. Another treatment strategy is to preserve existing photoreceptor function or survival by delivery of cytokines or growth factors to the diseased retina. In this case, we need to understand how the factors affect the photoreceptor cell (and other exposed cells as well). Since several of the factors being tested in clinical trials have key roles in photoreceptor development (and retinal development in general), our understanding of these processes is essential for evaluating the cause-and-effect relationships between the factors and the observed changes in treated photoreceptors. In a review article published in 2000, we and our postdoctoral mentor, Dr. Thomas Reh, described what was known at the time regarding the importance of cellular interactions and the identities and roles of soluble factors in rod photoreceptor development [1]. In this chapter, we summarize the state of the field, with particular attention paid to the findings made since 2000 and with an emphasis on both cell-intrinsic and -extrinsic factors.

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BRIEF OVERVIEW OF RETINAL DEVELOPMENT AND EARLY STAGES OF ROD PHOTORECEPTOR DIFFERENTIATION To appreciate the complexity of photoreceptor differentiation, it is important to have a basic understanding of how the retina develops. Because of space limitations, we can only briefly outline the major points. However, there are many excellent reviews that consider retinal development from various perspectives, and listed here is a very limited sampling of what is available [2–8]. The seven neuroretinal cell classes all derive from a common pool of progenitor cells known as retinal progenitor cells (RPCs). RPCs are resident in the optic neuroepithelium and are produced as a result of patterning events that occur during optic vesicle formation. Early on, the RPCs proliferate extensively and soon after initiate neuro- and gliogenesis. These processes, collectively termed retinal histogenesis, occur over significantly different developmental timescales among different vertebrate species, but certain key aspects of retinal histogenesis are remarkably well conserved. For example, there is a temporal progression in the production of cells in each class, and this order does not vary appreciably. Retinal ganglion cells, horizontal cells, and cone photoreceptors are born the earliest, followed by amacrine cells and rod photoreceptors; born last are the bipolar cells and Müller glia. As an example, Rapaport et al. [9] published an impressive quantitative analysis of retinal histogenesis in the rat. Cell marking-based lineage analyses revealed that as a population, RPCs are multipotential and lineage independent, meaning that these cells can produce more than one cell type, and their pattern of cell division is not deterministic with respect to cell output (although see [10] for a provocative contrast). An important issue then is to understand how the temporal order is established. Several now-classic cell culture and cell ablation experiments led to the model proposing that extracellular environmental factors are the primary driving forces behind retinal histogenesis and the regulation of its temporal progression. As a result, many extracellular signals and their signal transduction pathways are now known that influence histogenesis, and their precise functions and interactions are actively under investigation. More recent studies showed that RPCs are not completely naïve, however. A modification of the above model states that the responsiveness of RPCs to signals changes as development proceeds, and that this is regulated by changes in gene expression. It is proposed that these changes influence the competence of RPCs to produce specific cells and to bias their output in a temporal and progressive manner. Thus, a more complete model of retinal histogenesis incorporates both cell-extrinsic and -intrinsic factors, and much effort is currently devoted to understanding how these factors are integrated into specific networks. It is generally thought that cell-type-specific differentiation does not occur until the RPC exits the cell cycle. This is based in part on the outcomes of the lineage experiments (e.g., a single RPC division [two-cell clone]) can produce daughter cells with different fates) and on the collective observations that the expression of markers specific or deterministic for any single cell class or cell type is extremely rare in RPCs. Soon after RPCs exit the cell cycle, changes in cellular behavior and gene expression begin, and this continues until differentiation is complete. In the case of murine rod photoreceptors, this progression can last for over 2 weeks. Since the period of murine rod photoreceptor

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generation from RPCs is known and certain early morphological and molecular markers exist, postmitotic cells differentiating along the rod photoreceptor pathway can be defined as rod precursors, which distinguish them from RPCs. Rod precursors are also distinguishable from mature rod photoreceptors on the basis that rod inner and outer segment morphogenesis and synaptogenesis are barely under way, and the genes specific for rod phototransduction are not yet expressed. Rods have been traditionally identified as such when they express the rhodopsin gene because of its exquisite cell type specificity. Perhaps because of this, many studies of rod development have focused on the cellular parameters and molecular factors associated with rhodopsin expression. For example, the timing of activation of rhodopsin expression in the rat retina varies between rod precursors born before and those born after E19 [11, 12] and the timing of rhodopsin expression in other species has been correlated with the timing of cone opsin expression in neighboring cells (see Chapter 1). There are now several signaling pathways and transcription factors identified that regulate these parameters as well as control the expression of other factors important for rod function and survival. As another example, the exquisite cell type specificity of rhodopsin expression prompted efforts to identify the cis-acting DNA sequences and associated transcription factors that regulate rhodopsin expression. Many of those factors, perhaps not surprisingly in retrospect, play important roles in the pathway leading to mature rods. In the next sections, we describe those factors that in recent work were strongly implicated in controlling the important early steps in rod photoreceptor differentiation. Table 1 summarizes extracellular and transcription factors implicated in different aspects of rod photoreceptor development. Included in the table are those factors we are not able to describe due to space constraints. TRANSCRIPTION FACTORS Basic Helix-Loop-Helix Genes Basic helix-loop-helix (bHLH) genes constitute a large class of transcription factors that contain a stretch of positively charged amino acids termed the basic motif, which is important for DNA binding, and a helix-loop-helix domain, which is important for dimerization with other bHLH proteins. The bHLH genes have important roles in the nervous system, from the earliest stages of embryogenesis to the survival and function of mature neurons. In the Drosophila retina, the bHLH gene atonal has an essential role in specifying the R8 photoreceptor, which sets in motion the wave of neurogenesis in each presumptive ommatidial unit [35]. No fewer than ten bHLH genes have been identified in the mammalian retina, and most have some role either in RPC maintenance or in promoting cell fate and differentiation of neurons and Müller glia [36–38]. Among the retinal expressed bHLH genes, Neuro D may have the most direct role in photoreceptor development because of its abundant, although not necessarily exclusive, expression in photoreceptor precursors and because its over- or misexpression is sufficient to promote photoreceptor differentiation in RPCs or in nonneural ocular cells such as RPE and iris epithelial cells [39–47]. Neuro D expression is also correlated with de novo photoreceptor differentiation in damage-induced regeneration in the chick retina [48], and Neuro D is expressed in the rod precursor lineage in the adult goldfish retina [49], a

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Levine and Fuhrmann Factors regulating differentiation of rod photoreceptors

Factor Otx2 CRX QRX NRL NR2E3 Retinoic acid NeuroD Wnt4 Sonic hedgehog FGF Laminin β2 PEDF VEGF GDNF Activin

IGF-1 Rb Taurine

WIF-1 CNTF, LIF

EGFR

Description Activators TF, required for photoreceptor precursors TF, sufficient for photoreceptor fate, essential for expression of photoreceptor-specific genes (function) but not for fate TF, interacts with NRL and CRX, regulates survival or maturation of rods in chick and frog TF, sufficient and required for rod differentiation Nuclear receptor, suppresses cone fate, interacts with NRL to promote rod fate Nuclear receptor, stimulates rod differentiation, promotes NRL and CRX expression TF, promotes photoreceptor differentiation, required for rod differentiation in chick Secreted glycoprotein, promotes rod differentiation in vitro Morphogen, accelerates rod differentiation in vitro in rat, required for rod differentiation in zebrafish in vivo Fibroblast growth factor, promotes rhodopsin expression in vitro ECM component, promotes rod differentiation and required for inner and outer segment elongation Pigment epithelium-derived factor, promotes photoreceptor differentiation in frog Vascular endothelial growth factor, stimulates rod differentiation Glia-derived neurotrophic factor, promotes differentiation and survival of chick rods in vitro Growth factor, promotes rod differentiation in rat retinal cultures and inhibits terminal photoreceptor differentiation in chick Insulin-like growth factor, stimulates formation of rods in teleost fish Retinoblastoma protein, required for rod differentiation in mouse Promotes differentiation of rod photoreceptors in vivo and in vitro Inhibitors ECM component, inhibits rod differentiation in vitro Cytokines, inhibit terminal rod photoreceptor differentiation transiently in mammals via STAT3 phosphorylation and SOCS3 activation, can induce cell death of rod precursors Epidermal growth factor receptor and ligands such as EGF and TGFα inhibit rod differentiation

ECM extracellular matrix, TF transcription factor

Reference See text See text See text See text See text See text See text See text 13, 14 15–19 20–22 23 24 25 26, 27

28, 29 30, 31 See text

See text See text

32–34

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specialized neurogenic population that gives rise to new rods throughout the life of the organism. Resolving the requirement of Neuro D, and for that matter, other bHLH genes, in rod photoreceptor differentiation has been more difficult. Although work done in the chick retina suggests that Neuro D is required for photoreceptor development in that organism [50], the predominant photoreceptor phenotype in Neuro D knockout mice is decreased survival [42, 43]. One reason why Neuro D may not be required for rod photoreceptor differentiation in mice is because of redundancy or functional compensation by other bHLH proteins in its absence. To address this, Akagi et al. (2004) [51] developed multiple bHLH triple-knockout mice, and the most severe deficit in photoreceptor number was observed in the Mash1−/−, Math3−/−, Neuro D−/− triple-knockout mouse. However, rod photoreceptor development is still evident even in this combinatorial mutant. Furthermore, the high levels of photoreceptor apoptosis observed in this mutant leaves it unresolved whether elimination of these three genes causes a direct problem in photoreceptor specification or differentiation. Compounding the problem further, other bHLH genes are upregulated when certain bHLH genes are genetically inactivated, and there is ample evidence for functional redundancy among these genes. Otx2 Otx2 is a homeobox gene that is widely expressed in early embryos and becomes progressively restricted to the anterior portion of the neural tube. Consistent with its expression pattern, the Otx2 knockout mouse is embryonic lethal and lacks forebrain structures [52–54]. In the developing eye, Otx2 is expressed in the developing RPE and retina, and genetic studies in compound mutants of Otx2 and its paralog Otx1 showed that these genes are essential for RPE development [55–57]. Retinal Otx2 expression has been examined in several vertebrate species, and some differences in timing of activation have been reported (i.e., proliferating RPC vs. postmitotic precursors) [55–60]. In mice, Otx2 is expressed early in retinal development, but in postmitotic cells and in a pattern consistent with the generation of retinal neurons, most notably photoreceptors. Otx2 expression in the photoreceptors is transient and is ultimately expressed in the inner nuclear layer (INL) in presumptive bipolar cells, horizontal cells, and Müller glia [61]. Baas et al. (2000) [62] showed that Otx2 subcellular localization (nuclear vs. cytoplasmic) is developmentally regulated and cell type specific. To investigate the role of Otx2 in neural retinal development, Nishida et al. (2003) [61] generated conditional knockouts by crossing mice with a floxed Otx2 allele to mice with Cre recombinase expression controlled by the Crx promoter. Although some Otx2expressing cells persisted in these mice, the changes in retinal cell differentiation were dramatic. There was an almost-complete absence of rhodopsin-expressing cells and a major increase in the number of cells expressing markers of amacrine cells. The magnitude of these changes suggests that loss of Otx2 culminated in a change in fate from rods to amacrine cells, and this is consistent with Otx2 overexpression experiments. However, there is a large increase in apoptotic cells, and this could account in part for the loss in rods. Thus, although the effects of Otx2 inactivation on the retina are likely

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to be complex, these experiments demonstrated the essential nature of Otx2 for rod photoreceptor development or survival. Crx Crx is a homeobox gene that is closely related to Otx2. Initial reports showed that mutations in the human CRX gene cause autosomal dominant cone-rod dystrophy (CORD2) and late-onset retinitis pigmentosa [63, 64]. Subsequent studies showed that both dominant and recessive CRX mutations are also linked to Leber’s congenital amaurosis (LCA7), suggesting that CRX function is important for photoreceptor development in humans [65–71]. As described next, work done in animal models and in biochemical studies strongly supports this idea. Crx was discovered by its sequence similarity to other known homeobox genes [63, 72] and in a DNA–protein interaction screen (one-hybrid screen) using a retinal complementary DNA expression library and the Ret4 DNA sequence found in the rhodopsin promoter as bait [73]. Like Otx2, Crx expression begins early in retinal development [58, 60, 72, 74–78]. However, Crx follows Otx2 activation and is likely to be a direct transcriptional target of Otx2 in postmitotic rod and cone precursors [61]. Whereas Otx2 expression is downregulated in photoreceptors and remains strong in the INL in the mature retina, Crx remains highly expressed in the outer nuclear layer (ONL) and in a smaller subset of cells in the INL than Otx2. The importance of Crx in mammalian photoreceptor formation is now well established. Overexpression of Crx is sufficient to bias cells toward a photoreceptor fate in the embryonic retina and in adult iris epithelial cells [72, 79], and overexpression of a dominant negative form of Crx interferes with outer segment formation [72]. Consistent with these findings, genetic inactivation of Crx in mice causes developmental disruptions in outer segment formation and synaptogenesis as well as decreased expression of many phototransduction genes [80–83]. Crx acts as a direct transcriptional activator, and many studies showed Crx to be an integral component of multiprotein complexes that occupy the regulatory regions of several rod and cone photoreceptor genes [60, 72, 73, 75, 84–94]. Several interacting proteins have also been found that interfere with Crx-mediated transactivation, and these interactions may be important for the timing of photoreceptor gene activation during differentiation. These interactions may also function to restrict the transcriptional activation function of Crx to rods or bipolar cells [95–99]. Crx has also been shown to directly interact with Ataxin-7, a protein that undergoes polyglutamine expansion and is responsible for spinocerebellar ataxia type 7 (SCA7). While the interaction with wild-type Ataxin-7 is important for the assembly of a Crx transactivation complex, the polyglutamine-expanded, neurodegenerative form of Ataxin-7 interferes with this interaction, resulting in a downregulation of photoreceptor gene expression, and this appears to be a causative factor for the cone-rod dystrophy found in SCA7 [100–103]. Interestingly, Crx is not essential for the initial specification and differentiation of photoreceptors, as evidenced by their appearance in the Crx−/− mouse retina. This is somewhat surprising given its early onset of expression. However, Bibb et al. (2001) [74] showed that, in the fetal human retina, the expression of several photoreceptor genes that

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are targets of Crx regulation in the mouse precedes the onset of CRX expression, which further suggests that Crx is not required for early photoreceptor development. Why cell fate specification and early differentiation of photoreceptors occur in the absence of Crx is not known. One possibility is that Otx2 partially compensates or is redundant with Crx. Consistent with this idea, Crx and Otx2 are both in the Otx gene family [58, 76, 104], both are expressed early in photoreceptor differentiation, Crx expression is not maintained in the Otx2−/− retina, and Otx2−/− mice show no evidence of photoreceptor formation [61]. Furthermore, Crx can interact with Mitf and promote pigment cell differentiation in vitro to an extent comparable to Otx1 and Otx2 [58]. However, experiments in Xenopus suggest that Otx5b (Crx) and Otx2 are not redundant in retinal cell type specification: Otx5b and Otx2 promote photoreceptor and bipolar cell fates, respectively [60]. Zebrafish contain two Crx-related genes (Crx and Otx5), and each seems to have unique functions in regulating gene expression in the retina and pineal gland, which in mammals appear to be relegated to Crx alone (mammals contain a single Crx/Otx5 gene) [59, 75, 105]. In contrast to other species, zebrafish Crx is also expressed in RPCs, and morpholino-mediated RNA knockdown disrupts proximo-distal patterning of the retina and causes a general delay in neuronal differentiation [59]. To ultimately address the issue of whether redundancy or compensation accounts for the nonessential nature of Crx in photoreceptor cell fate specification in the mouse retina, it may be necessary to create a dominant negative Crx allele that interferes with the functions of both Otx2 and Crx. QRX/Rax-L/Rx-L QRX is a homeobox gene in the Rx family. Rx proteins contain paired-like homeodomains and are distantly related to other important retinal transcription factors, such as Chx10, Vsx1, Pax6, and the Otx cohort. QRX is found in the human and bovine genomes [106], and putative orthologs are identified in chick (Rax-L) [107, 108] and Xenopus (Rx-L) [109]. The role of QRX/Rax-L/Rx-L is not as well established as the other genes described in this chapter for two reasons: An ortholog has not been identified in the mouse, and a congenital photoreceptor disease has not been found in humans. Nevertheless, QRX, Rax-L, and Rx-L have all been shown to bind to conserved DNA sequences in the rhodopsin promoter, interact with Nrl and Crx, and promote transcription [106, 107, 109]. Furthermore, dominant negative experiments in chick and gene knockdown experiments in Xenopus both supported a role in the maturation or survival of photoreceptors [107, 109]. NRL The Nrl gene is a member of the Maf transcription factor family. Maf proteins contain a basic motif linked to a leucine zipper domain (bZIP), which is required for DNA binding and protein dimerization, respectively [110]. Several Maf genes have important roles in lens development [110–114] and Nrl has emerged as a key player in rod photoreceptor development. Genetic inactivation of Nrl (Nrl−/−) leads to a complete loss of rods, but interestingly, their absence is not due to a failure in photoreceptor cell production or in decreased survival. Rather, the ONL in the Nrl−/− retina is well

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Fig. 1. Interaction between extracellular signals and transcription factors that results in a stepwise progression of rod photoreceptor development. See Table 1 for definitions of abbreviations.

populated, and these cells have the characteristic expression profiles and physiological and morphological properties of cones, in particular S cones [115–117]. These observations strongly suggest that Nrl is an essential transcription factor for executing the rod differentiation program and, at the same time, for keeping the cone differentiation suppressed. Consistent with this, many studies showed that Nrl is a direct positive regulator of rhodopsin gene transcription. A prediction of this model is that Nrl should be expressed in postmitotic photoreceptor precursors. While true, some studies suggested that Nrl is expressed in the inner retina, and its onset of expression is earlier than the period of rod genesis [118, 119]. However, more recent studies examining Nrl protein expression in human and mouse embryos [120, 121] and in a transgenic mouse line expressing green fluorescent protein (GFP) under the control of the Nrl promoter [122] suggested that Nrl may be specific to postmitotic rod precursors. Although the discrepancies in the reported Nrl expression characteristics are not completely resolved, Oh et al. (2007) expressed Nrl under the control of the Crx promoter in both Nrl+/+ and Nrl−/− mice [123]. In both genetic backgrounds, the expression of Nrl drove rod photoreceptor development and did so at the expense of cones. In addition, transgenic Nrl expression in the Nrl−/− retina rescued the dysmorphic architecture of the ONL. These dramatic results showed not only that Nrl is sufficient to drive rod photoreceptor development when expressed in early postmitotic photoreceptor precursors, but also that cone precursors are competent to execute a rod photoreceptor program. Thus, “photoreceptor precursor” may be a more accurate description of earlystage rod and cone precursors, and onset of Nrl expression may signify a transition to a more restricted rod precursor state (see Fig. 1 and Conclusions).

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Given its key function in mouse rod development, it is reasonable to question whether Nrl is of similar importance in the human retina. Interestingly, most of the identified human mutations in NRL are associated with adRP [124–127]. However, Nishiguchi et al. (2004) [128] recently reported two mutations that are likely to encode loss-of-function alleles, and patients with these mutations have clumped pigmentary retinal degeneration and some preservation of blue cone function, both of which are indicative of an analogous function for NRL in human photoreceptor differentiation as in mice. Nuclear Receptors Nuclear receptors constitute a large family of transcription factors that function as activators on direct binding of a ligand [129] (see Pharmacological Reviews, Volume 58, Issue 4, for an extensive overview). Those in which a physiological ligand is not known or the receptor is suspected to lack a ligand are listed as “orphan receptors.” Most nuclear receptors homodimerize or heterodimerize in various combinations and are associated with large multisubunit complexes that can regulate transcription in positive and negative manners. Because of their diversity in structure and function and their direct regulation by environmental cues, nuclear receptors influence a large number of cellular responses and gene expression programs. Several nuclear receptors are implicated in photoreceptor development; they include the thyroid hormone receptor beta 2 (TRβ2), several retinoid X receptors (RXRs) and retinoic acid receptors (RARs), and the orphan receptors Nr2E3 (PNR, RNR), and retinoid orphan receptor beta (RORβ). TRβ2, RXRγ, and RORβ are implicated in regulating the timing and selection of opsin expression in developing cones [91, 130–134]. Because of space limitations, however, we focus here on the proposed roles of nuclear receptors on rod development, and much of the current effort has been aimed at understanding the role of Nr2E3 in this process. Nr2E3 Patients with enhanced S-cone syndrome (ESCS) suffer from progressive vision loss that is initially characterized by night blindness and variability in L and M cone- (L/M cone-) mediated vision and ultimately followed by rod photoreceptor degeneration. What sets this disease apart from most other forms of inherited retinal degenerations is an increased sensitivity to blue light, which is mediated by the S cones. Jacobson and colleagues first described this condition in the early 1990s and noted the similarities in severely affected patients to those with Goldman-Favre syndrome [135–137]. Subsequent psychophysical and electrophysiological studies led to the prediction that patients with ECSC had an unusually high number of S cones, possibly at the expense of the other cones and rod photoreceptors [138–140]. It is now known that mutations in the NR2E3 gene can cause ESCS, Goldman-Favre syndrome, and clumped pigmentary retinal degeneration, and histopathological studies confirmed an overabundance of S-opsin-expressing cells and a lack of rods [141–145]. Expression analysis of NR2E3 during human fetal development showed the transcript and protein to be expressed in a pattern most consistent with postmitotic differentiating rod precursors [146]. rd7 is a naturally derived mutation in Nr2E3, and it serves as an excellent model for the human NR2E3 mutation [141, 147]. As in humans, the rd7 allele is a recessive

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allele, although the exact nature of the mutation was not clear until recently [148]. Nevertheless, the long-standing interpretation that the rd7 mutation is a null still holds, and homozygotes have the ECSC phenotype and gradual photoreceptor degeneration. It is also well established that the enhanced S-cone activity is due primarily to changes that occur during retinal development. However, two models exist to explain how Nr2E3 functions in photoreceptor development; they are described next. One model states that Nr2E3 functions to prevent the excess production of cone photo receptors from RPCs by restricting their proliferation after the normal interval of cone photoreceptor generation [149, 150]. This model makes several predictions. One prediction is that a subset of late embryonic RPCs is committed, or at the very least, specified, to give rise to cones and is actively suppressed from doing so by Nr2E3. Although there is no evidence for a cone-restricted RPC from lineage studies in mice [151], ectopic proliferation is observed in the rd7 retina and it is suggested that these proliferative cells are either S-cone precursor cells that are normally postmitotic or are RPCs fated to give rise to cones [149, 150]. Another prediction is that Nr2E3 is expressed in RPCs after the normal period of cone production. While one study reports Nr2E3 expression in proliferating RPCs at the expected time [149], several others argued that Nr2E3 expression is initiated in postmitotic rod precursors [146, 152, 153], which are also generated during this interval. Expression analyses in zebrafish and mouse suggested that Nr2E3 is transiently expressed in postmitotic immature cones [152, 153], although one study argued that Nr2E3 expression in cones is not transient [149]. Another prediction is that any role for Nr2E3 in rod development is independent of its role in restricting cone production. The evidence given for this is that the size of the S-cone cell population in the rd7 retina does not increase at the expense of L/M cones or rods (the reduction in rods is proposed to occur by apoptosis [149]). While there is general agreement that the L/M cone population is not affected in the rd7 retina, there is disagreement on whether the extra S-cone cells originate from a source of ectopically proliferating cells or from cells normally fated to become rods, which brings us to the next model. The second model states that Nr2E3 functions as a differentiation factor for postmitotic rod precursors and does this in two ways: by repressing the cone gene expression program by acting as a transcriptional repressor and by facilitating the rod gene expression program by acting as a transcriptional activator. The evidence for this model is compelling on several levels. First, regardless of whether Nr2E3 is expressed in RPCs or cones, it is clear that it is expressed in postmitotic rod photoreceptors. Second, in the rd7 retina, many cone messenger RNAs (mRNAs) are highly upregulated and expressed throughout the ONL, which is largely composed of rods as determined by the expression of rod genes [149, 152–154] and by the presence of initially normal scotopic electroretinogram (ERG) responses [147]. Third, several studies have documented that Nr2E3 can function as a transcriptional repressor or activator, and this depends on the context of the cis-acting DNA regulatory sequences and on the protein complexes with which Nr2E3 is associated [150, 152, 153, 155]. It is important to note, however, that many rod genes are not significantly downregulated in the rd7 retina, and this could be due to the persistent expression of Crx, Nrl, and other photoreceptor transcription factors [149, 152–155]. Along these lines, comparison of the Nrl−/− and rd7 photoreceptor phenotypes shows considerable overlap, especially with respect to increases in the number of photoreceptors with

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S-cone gene expression characteristics. One difference, however, is that in the Nrl−/− retina, the expression of rhodopsin and other rod-specific phototransduction genes is significantly downregulated. Correlated with this is a lack of Nr2E3 expression [116]. Cheng et al. (2006) [156] developed transgenic mice expressing Nr2E3 from the Crx promoter and S-opsin promoter and analyzed the photoreceptor phenotypes in normal and Nrl−/− mice. Nr2E3 is activated in early-stage rod and cone precursors (photoreceptor precursor stage) when driven by the Crx promoter and is activated in S-cone precursors by the S-opsin promoter. The primary findings of this work are that Nr2E3 is sufficient to block cone gene expression in the presence or absence of Nrl, and it can activate rod gene expression in the absence of Nrl, although not completely. The sum of these observations supports the idea that Nr2E3 suppresses the expression of the cone phenotype, at the same time promoting the rod phenotype in combination with Nrl. Are these models mutually exclusive? While certain aspects of these models are discordant and need to be resolved, it is possible that both could be correct in their essential aspects. It is important to note that not all photoreceptors in the rd7 mouse are functional cones. In fact, it is estimated that the increase in S-cone number is from approximately 1% to 3% of the total retinal cells [154, 157]. Second, although it is clear that cone mRNA expression is inappropriately activated in the rd7 rod population, cone protein expression appears to be limited [149] and S-opsin protein has not been found in rhodopsin-expressing cells [154]. While the evidence is strong that Nr2E3 has a role in suppressing the cone and promoting the rod gene expression programs in developing rods, the source of the supernumary S cones, whether from cells that underwent ectopic proliferation or from cells normally fated to be rods, still needs to be resolved. Retinoic Acid/Retinoic Acid Receptors Vitamin A derivatives (retinoids) regulate many aspects of vertebrate development and homeostasis. They bind to and activate nuclear receptor proteins of the steroid and thyroid hormone receptor superfamily, resulting in derepression and transcriptional activation of target genes. Homo- or heterodimers of RARs and RXRs are activated by retinoids with high affinity, whereas unsaturated fatty acids such as docosahexaenoic or linoleic acids function as low-affinity ligands. Retinoic acid-synthesizing enzymes and receptors are present in the vertebrate eye during photoreceptor development [131, 158–162]. Several decades ago, Dowling observed that vitamin A is essential for development and maintenance of photoreceptors in cats [163]. Culture experiments in chick and rodents showed subsequently that the primary metabolite retinoic acid stimulates photoreceptor differentiation and survival [164–166]. In zebrafish, retinoic acid treatment resulted in precocious rod differentiation, whereas inhibition of synthesis using citral led to a delay in photoreceptor differentiation [167]. Similarly, retinoic acid injections into pregnant rats late during pregnancy accelerated rod differentiation after birth [168]. Another ligand for RXRs, docosahexaenoic acid (DHA), is a major structural lipid of retinal photoreceptor outer segment membranes. In rat retinal cultures, DHA promotes maturation of CRX-positive photoreceptor precursors [169]. Thus, these studies indicate that RAR signaling is sufficient and required for photoreceptor differentiation in vertebrates.

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At which stage does retinoic acid act to regulate rod development? In vitro, double-labeling experiments with BrDU and rod markers indicated that retinoic acid acts on proliferating RPCs to increase the proportion of rods. This effect was accompanied by a decrease of amacrine cells, suggesting that retinoic acid instructs the rod fate in RPCs by compromising the generation of other cell types [164]. Conversely, in zebrafish, treatment with retinoic acid increased rhodopsin expression, but did not alter cell numbers, arguing against an effect on cell fate [170]. However, Khanna et al. showed that retinoic acid can directly transactivate the Nrl promoter and promotes expression of NRL in mammalian retinal cells in vitro [171]. Thus, retinoic acid indeed could act as an early extrinsic signal to instruct RPCs to adopt the rod fate. In agreement with this, Li et al. observed an increase of CRX expression in retinoic acid-treated human retinoblastoma cells [172]. In sum, these observations suggest that the outcome of retinoic acid action on rod development is species dependent. Interestingly, in mouse retinal explants physiological concentrations of retinoic acid have also been shown to induce rod-specific cell death when RPE is present [173]. Furthermore, since retinoic acid can induce expression of cone and rod photoreceptorspecific genes in retinoblastoma cell lines [172, 174–176], it might exert differential effects that activate distinct mechanisms in both cell types: alone to regulate rod differentiation or in combination with TRβ2 to regulate cone differentiation [131].

EXTRACELLULAR FACTORS AND SIGNAL TRANSDUCTION PATHWAYS Wnt/Frizzled Pathway Wnts are secreted glycoproteins that bind to Frizzled transmembrane receptors and regulate multiple developmental processes such as embryonic patterning, cell polarity, cell fate determination, and proliferation. Many Wnts, Frizzleds, and pathway modulators are expressed during retinal development, and more recently, Wnt/Frizzled signaling has been shown to regulate several aspects of eye development in vertebrates [177, 178]. Wnt-inhibitory factor (WIF-1) is a secreted antagonist of the Wnt/b-catenin pathway that is present in the extracellular matrix (ECM) and binds to Wnts in the extracellular space [179, 180]. It contains a signal sequence, a WIF domain, and five epidermal growth factor (EGF) repeats. Hunter et al. (2004) [180] identified WIF as an ECM component in the developing mouse retina. During the period of photoreceptor differentiation, WIF-1 is expressed in the subretinal space and interphotoreceptor matrix. Other potential partners of the Wnt pathway that are expressed appropriately are Wnt4, LRP6, and Frizzled-4. Furthermore, WIF-1 and Wnt4 appear to interact with each other [180]. In rat retina dissociated cultures, WIF-1 decreases, and Wnt4 promotes rod differentiation, suggesting that these components could modulate rod differentiation and maintenance. However, Frizzled-4 mutants do not show a photoreceptor phenotype, and Wnt/β-catenin activity in the photoreceptor layer of postnatal mice is not detected in Wnt/reporter mice [181, 182] (our unpublished observations). Furthermore, conditional inactivation of b-catenin in the mouse does not result in an obvious change of rhodopsin immunoreactivity [183].

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These studies suggest Wnt/β-catenin signaling might not be active and required for differentiation of postnatal photoreceptors. Taurine Another positive regulator of rod differentiation is taurine, a cystein derivative that is structurally similar to γ-aminobutyric acid (GABA) and glycine. It is highly expressed in the developing and mature retina. When omitted in the diet, developmental defects of brain and retina occur in kittens (for review, see [184]). Moreover, Altshuler et al. showed that taurine promotes rod differentiation in rat retinal cultures [185]. Interestingly, taurine can activate glycine and GABA receptors, and the action of taurine is dependent on the activity of these receptors [184]. In mouse retinal cultures, application of glycine or GABA receptor antagonists abolished the effects of taurine on rod differentiation. Importantly, inhibition experiments using small interfering RNA (siRNA) against glycine receptor a 2 (GlyRa 2) revealed that fewer photoreceptors develop in vivo, which is compensated by a higher number of late-born cell types such as Müller glia and bipolar cells [184]. These and other experiments suggest that taurine/ glycine receptor signaling negatively regulates RPC proliferation and stimulates rod differentiation. This effect appears to be specific for the photoreceptor lineage since forced expression for GlyRα2 can induce photoreceptor-specific genes in E16 retinal progenitors. Conversely, without taurine/glycine receptor activation, progenitor cells might keep dividing and differentiate eventually into late-born cell types due changes in the extracellular environment. In agreement with the role of taurine and GlyRα2 during central nervous system development, disruption of the GlyRa 2 gene results in defects of electrophysiological and calcium imaging responses of cortical cells [186]. However, no abnormalities in development and function of retinae of GlyRa 2−/− could be observed; Nrl and rhodopsin expression and ERG analysis are normal without GlyRα2 [186]. Since expression of the other glycine receptor subunits is not changed, these results suggest that GABA receptor signaling may compensate or glycine receptor activation is not essential for rod development. Ciliary Neurotrophic Factor/Leukemia Inhibitory Factor/Pleiotrophin/Signal Transducer and Activators of Transcription 3/SOCS Ciliary neurotrophic factor (CNTF) is a member of the interleukin 6 (IL-6) cytokine family and activates a trimeric transmembrane receptor complex consisting of the shared β receptors gp130 and LIFRβ (leukemia inhibitory factor receptor β) and a CNTFRαspecific subunit. Ligand binding leads to activation of gp130 and subsequently of associated tyrosine kinases (Janus kinases; JAKs), which phosphorylate the β subunits and signal transducer and activators of transcription (STAT) 1 and 3. CNTF can activate three pathways: JAK/STAT, mitogen-activated protein (MAP) kinase, and Akt/PI3 K signaling [187–191]. CNTF, other CNTF-like ligands, and LIF (activation by dimerization of gp130 and LIRFβ) exert diverse effects on photoreceptors in vertebrates. In developing chick photo receptors, CNTF promotes expression and relocalization of opsin proteins in cultured photoreceptors, as shown by labeling with the rho4D2 antibody, which recognizes

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rhodopsin and green cone opsin [192–194]. Although no such effect of CNTF was found on cones using peanut lectin as a marker [192], Adler’s laboratory later observed that CNTF specifically promotes expression of green cone opsin but not rhodopsin and, interestingly, can induce coexpression of green and red cone opsin [195, 196]. Thus, in the chick these experiments indicated that CNTF produced by Müller glia might function to increase production of green cone opsin specifically, which is supported by the expression pattern of CNTFRα in the photoreceptor layer [197–199]. However, loss-of-function experiments need to be performed to obtain further insight of a regulatory role of CNTF in developing chick photoreceptors in vivo. In the adult mammalian retina, CNTF protects, most likely indirectly, photoreceptors in models of retinal degeneration and injury [200–207]. In the developing rodent retina, CNTF exerts a different role. Here, CNTFR signaling controls the timing of terminal differentiation and the number of rod photoreceptors by acting on photoreceptor precursors. These functions are generally in good agreement with the spatiotemporal expression pattern of LIF, CNTF, and CNTF-like ligands such as neuropoeitin, CLF/ CLC rodents [208–210], or possible downstream mediators such as pleiotrophin [211], receptor expression [212–216], and with the activation of the signal transduction machinery [215, 217–219]. First shown in dissociated or explant cultures, CNTF and LIF act to suppress recoverin and rhodopsin expression [194, 220–223]. Interestingly, CNTF treatment does not alter the morphogenesis of rod outer segments; it acts reversibly on rod progenitors [218, 223]. The CNTF effect is transient since photoreceptor precursors become less responsive to CNTF with increasing age, and one explanation could be that CNTF receptor expression in photoreceptors decreases with maturation [218, 219, 221, 223]. These observations are supported by the timing of expression and activation of CNTFR pathway components in vivo and suggest that CNTF acts as a negative regulator pre- and perinatally to delay terminal photoreceptor differentiation [194, 215]. It is possible that formation of outer segments and concomitant changes in the surrounding ECM might interfere with cell division in the apical layer necessary to produce bipolar and Müller glia [223, 224]. Thus, the long delay between birth of photoreceptor precursors and their terminal differentiation could function to enable proper maturation of late-born retinal cell types. At higher CNTF concentrations (e.g., 20 ng/ml), more cells with bipolar-specific gene expression appear in vitro [220, 223, 225, 226]. Originally proposed as an effect on switch of cell fate, in vivo and in vitro studies now indicate that CNTF and LIF keep photoreceptor precursors in an immature state without affecting specification of other cell types [209, 220–223, 227]. For example, disruption of the CNTFRα gene leads to an increase or acceleration of rod differentiation in explant cultures but does not affect bipolar cells [220]. Very similar observations were made following in vivo injection of an inhibitor of the CNTF/LIF pathway [209]. A possible explanation is that CNTF might upregulate expression of its own receptor in vitro, suggesting that the promoting effect on bipolar cells depends on culture conditions and does not reflect a physiological role in vivo [223]. Although capable of activating different pathways, the CNTF effects on rod development are exclusively mediated through rapid phosphorylation and activation of STAT3, which is independent of EGF signaling, another inhibitor of rod differentiation

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[217–219]. CNTF appears to act on postmitotic rod precursors that express CRX and on newly differentiating rhodopsin-positive photoreceptors directly by increasing STAT3 phosphorylation in these cells [217–219]. However, phosphorylated STAT3 might not directly regulate rhodopsin transcription but rather upstream regulators. Supporting evidence comes from experiments using overproduction of LIF in transgenic mice [227]. Strong activation of LIF signaling early during retinal development results in suppressed expression of factors regulating the photoreceptor lineage, commitment to the rod fate, and rhodopsin production, such as CRX, Neuro D and both Nrl and Nr2E, which are required to transactivate the rhodopsin promoter. Interestingly, also inhibitory effects on cone-specific genes such as TRβ2 and S- and M-cone opsin were observed [227]. A new regulatory mechanism has been described that may function to ensure the maturation of rods in an otherwise inhibitory environment [218]. Residual STAT3 activation in the postnatal rodent retina is suppressed by upregulation of an intracellular negativefeedback modulator: suppressor of cytokine signaling 3 (SOCS3). SOCS3 expression is induced by cytokine signaling and inhibits STAT3 phosphorylation by blocking JAK activity. Conditional inactivation of SOCS3 delays photoreceptor differentiation and results in decreased CRX expression, whereas SOCS3 overexpression makes photoreceptor precursors insensitive to CNTF [228]. Interestingly, CNTF has also been proposed to function in regulating programmed cell death of postnatal photoreceptors [209]. In the postnatal mouse retina, natural cell death of photoreceptor precursor cells occurs, possibly as a correction mechanism to limit the number of differentiated photoreceptors. These cells are called “inner rods” because they are observed in the outer INL [229]. Elliott et al. (2006) [209] showed that inhibition of CNTFR/LIFR signaling or nitric oxide synthase activity in vivo decreases apoptotic cell death in this specific population. This effect was observed quickly after treatment (20 h), suggesting a direct effect on photoreceptor survival. Similarly, Graham et al. observed that overexpression of LIF leads to reduced thickness of the photoreceptor layer in the postnatal retina, although they did not investigate whether cell death is increased [227]. Since CNTF does not contain a signal sequence, secreted CNTF-like ligands such as LIF or CLC/CLF are likely candidates; they also increased developmental cell death in the rodent retina in vivo [209]. Together, these studies indicate that a more precise understanding of the mechanism of CNTF action is necessary, especially since CNTF is one of the factors currently being tested in clinical trials to prevent photoreceptor cell death in retinal degeneration. CONCLUSIONS AND FUTURE PROSPECTS In Fig. 1, we propose a model of how the factors we discussed here may fit together to regulate rod photoreceptor differentiation. As RPCs exit the cell cycle, they begin to express transcription factors that in part define a precursor cell of photoreceptors, both rod and cone. As the photoreceptor precursor develops, it begins to express new transcription factors and respond to signals that push it into a more restricted fate, such as the rod precursor, and ultimately into a rod photoreceptor. While we hope to convey here the extraordinary progress made thus far in understanding rod photoreceptor development, we would argue that we still have a relatively

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rudimentary understanding of the cellular and molecular mechanisms. For example, it is still not clear when and if a photoreceptor precursor becomes irreversibly committed to becoming a rod. And, if commitment does occur, what are the necessary factors to ensure this? Photoreceptor precursors show a high degree of morphological plasticity [230] and molecular plasticity, and it is not clear whether this reflects an uncommitted precursor or a committed precursor passing through steps of maturation. As Adler lucidly described in a recent review [3], the study of cell commitment in the retina is extremely difficult and requires further development of tools to understand this process from a prospective outlook. However, progress is being made. MacLaren et al. (2006) [231] showed in an elegant study that Nrl expression in photoreceptor precursors significantly enhances the potential of these cells to differentiate into rods and to integrate into normal or degenerating retina when transplanted. This is an important advance with potential clinical implications because it suggests that Nrl may function as a rod commitment factor, even at a time when these cells are morphologically immature. Thus, even though several potential stem cell sources can be manipulated to produce photoreceptors [45, 232–242], this typically occurs at low frequencies, and it will be important to determine if Nrl expression can be more efficiently induced and, if so, if this is sufficient for robust rod production. Another important area of research that should make a significant stride in the next few years addresses how later aspects of photoreceptor differentiation are regulated, such as the formation of apical-basal polarity and assembly of the inner and outer segments. In this regard, mutations from genetic screens in zebrafish and Drosophila are beginning to provide some initial insights [243, 244]. It will be an especially gratifying day when we will have a comprehensive and integrated model of photoreceptor development from the earliest cell fate decisions in the RPC to the final assembly of the outer segment in the mature rod. REFERENCES 1. Levine, E. M., Fuhrmann, S., Reh, T. A. (2000). Soluble factors and the development of rod photoreceptors. Cell Mol Life Sci 57, 224–234. 2. Livesey, F. J., Cepko, C. L. (2001). Vertebrate neural cell-fate determination: lessons from the retina. Nat Rev Neurosci 2, 109–118. 3. Adler, R. (2005). Challenges in the study of neuronal differentiation: a view from the embryonic eye. Dev Dyn 234, 454–463. 4. Chow, R. L., Lang, R. A. (2001). Early eye development in vertebrates. Annu Rev Cell Dev Biol 17, 255–296. 5. Fuhrmann, S., Chow, L., Reh, T. A. (2000). Molecular control of cell diversification in the vertebrate retina. In: Vertebrate eye development (Fini, M. E., ed.), Vol. 31, pp. 69–91. Springer, Berlin. 6. Reh, T. A., Levine, E. M. (1998). Multipotential stem cells and progenitors in the vertebrate retina. J Neurobiol 36, 206–220. 7. Zhang, S. S., Fu, X. Y., Barnstable, C. J. (2002). Molecular aspects of vertebrate retinal development. Mol Neurobiol 26, 137–152. 8. Yang, X. J. (2004). Roles of cell-extrinsic growth factors in vertebrate eye pattern formation and retinogenesis. Semin Cell Dev Biol 15, 91–103.

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9. Rapaport, D. H., Wong, L. L., Wood, E. D., Yasumura, D., LaVail, M. M. (2004). Timing and topography of cell genesis in the rat retina. J Comp Neurol 474, 304–324. 10. Cayouette, M., Barres, B. A., Raff, M. (2003). Importance of intrinsic mechanisms in cell fate decisions in the developing rat retina. Neuron 40, 897–904. 11. Belliveau, M. J., Young, T. L., Cepko, C. L. (2000). Late retinal progenitor cells show intrinsic limitations in the production of cell types and the kinetics of opsin synthesis. J Neurosci 20, 2247–2254. 12. Morrow, E. M., Belliveau, M. J., Cepko, C. L. (1998). Two phases of rod photoreceptor differentiation during rat retinal development. J Neurosci 18, 3738–3748. 13. Levine, E. M., Roelink, H., Turner, J., Reh, T. A. (1997). Sonic hedgehog promotes rod photoreceptor differentiation in mammalian retinal cells in vitro. J Neurosci 17, 6277–6288. 14. Stenkamp, D. L., Frey, R. A., Prabhudesai, S. N., Raymond, P. A. (2000). Function for Hedgehog genes in zebrafish retinal development. Dev Biol 220, 238–252. 15. Hicks, D., Courtois, Y. (1988). Acidic fibroblast growth factor stimulates opsin levels in retinal photoreceptor cells in vitro. FEBS Lett 234, 475–479. 16. Hicks, D., Courtois, Y. (1992). Fibroblast growth factor stimulates photoreceptor differentiation in vitro. J Neurosci 12, 2022–2033. 17. McFarlane, S., Zuber, M. E., Holt, C. E. (1998). A role for the fibroblast growth factor receptor in cell fate decisions in the developing vertebrate retina. Development 125, 3967–3975. 18. Patel, A., McFarlane, S. (2000). Overexpression of FGF-2 alters cell fate specification in the developing retina of Xenopus laevis. Dev Biol 222, 170–180. 19. Zhang, L., El-Hodiri, H. M., Ma, H. F., Zhang, X., Servetnick, M., Wensel, T. G., Jamrich, M. (2003). Targeted expression of the dominant-negative FGFR4a in the eye using Xrx1A regulatory sequences interferes with normal retinal development. Development 130, 4177–4186. 20. Hunter, D. D., Brunken, W. J. (1997). Beta 2 laminins modulate neuronal phenotype in the rat retina. Mol Cell Neurosci 10, 7–15. 21. Hunter, D. D., Llinas, R., Ard, M., Merlie, J. P., Sanes, J. R. (1992). Expression of s-laminin and laminin in the developing rat central nervous system. J Comp Neurol 323, 238–251. 22. Libby, R. T., Hunter, D. D., Brunken, W. J. (1996). Developmental expression of laminin beta 2 in rat retina. Further support for a role in rod morphogenesis. Invest Ophthalmol Vis Sci 37, 1651–1661. 23. Jablonski, M. M., Tombran-Tink, J., Mrazek, D. A., Iannaccone, A. (2000). Pigment epithelium-derived factor supports normal development of photoreceptor neurons and opsin expression after retinal pigment epithelium removal. J Neurosci 20, 7149–7157. 24. Yourey, P. A., Gohari, S., Su, J. L., Alderson, R. F. (2000). Vascular endothelial cell growth factors promote the in vitro development of rat photoreceptor cells. J Neurosci 20, 6781– 6788. 25. Rothermel, A., Layer, P. G. (2003). GDNF regulates chicken rod photoreceptor development and survival in reaggregated histotypic retinal spheres. Invest Ophthalmol Vis Sci 44, 2221–2228. 26. Belecky-Adams, T. L., Scheurer, D., Adler, R. (1999). Activin family members in the developing chick retina: expression patterns, protein distribution, and in vitro effects. Dev Biol 210, 107–123. 27. Davis, A. A., Matzuk, M. M., Reh, T. A. (2000). Activin A promotes progenitor differentiation into photoreceptors in rodent retina. Mol Cell Neurosci 15, 11–21. 28. Mack, A. F., Fernald, R. D. (1993). Regulation of cell division and rod differentiation in the teleost retina. Brain Res Dev Brain Res 76, 183–187.

52

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29. Zygar, C. A., Colbert, S., Yang, D., Fernald, R. D. (2005). IGF-1 produced by cone photoreceptors regulates rod progenitor proliferation in the teleost retina. Brain Res Dev Brain Res 154, 91–100. 30. Zhang, J., Gray, J., Wu, L., Leone, G., Rowan, S., Cepko, C. L., Zhu, X., Craft, C. M., Dyer, M. A. (2004). Rb regulates proliferation and rod photoreceptor development in the mouse retina. Nat Genet 36, 351–360. 31. Johnson, D. A., Donovan, S. L., Dyer, M. A. (2006). Mosaic deletion of Rb arrests rod differentiation and stimulates ectopic synaptogenesis in the mouse retina. J Comp Neurol 498, 112–128. 32. Lillien, L., Cepko, C. (1992). Control of proliferation in the retina: temporal changes in responsiveness to FGF and TGF alpha. Development 115, 253–266. 33. Reh, T. A. (1992). Cellular interactions determine neuronal phenotypes in rodent retinal cultures. J Neurobiol 23, 1067–1083. 34. Lillien, L. (1995). Changes in retinal cell fate induced by overexpression of EGF receptor. Nature 377, 158–162. 35. Hsiung, F., Moses, K. (2002). Retinal development in Drosophila: specifying the first neuron. Hum Mol Genet 11, 1207–1214. 36. Hatakeyama, J., Kageyama, R. (2004). Retinal cell fate determination and bHLH factors. Semin Cell Dev Biol 15, 83–89. 37. Vetter, M. L., Brown, N. L. (2001). The role of basic helix-loop-helix genes in vertebrate retinogenesis. Semin Cell Dev Biol 12, 491–498. 38. Yan, R. T., Ma, W., Liang, L., Wang, S. Z. (2005). bHLH genes and retinal cell fate specification. Mol Neurobiol 32, 157–171. 39. Acharya, H. R., Dooley, C. M., Thoreson, W. B., Ahmad, I. (1997). cDNA cloning and expression analysis of NeuroD mRNA in human retina. Biochem Biophys Res Commun 233, 459–463. 40. Ahmad, I., Acharya, H. R., Rogers, J. A., Shibata, A., Smithgall, T. E., Dooley, C. M. (1998). The role of NeuroD as a differentiation factor in the mammalian retina. J Mol Neurosci 11, 165–178. 41. Kanekar, S., Perron, M., Dorsky, R., Harris, W. A., Jan, L. Y., Jan, Y. N., Vetter, M. L. (1997). Xath5 participates in a network of bHLH genes in the developing Xenopus retina. Neuron 19, 981–994. 42. Morrow, E. M., Furukawa, T., Lee, J. E., Cepko, C. L. (1999). NeuroD regulates multiple functions in the developing neural retina in rodent. Development 126, 23–36. 43. Pennesi, M. E., Cho, J. H., Yang, Z., Wu, S. H., Zhang, J., Wu, S. M., Tsai, M. J. (2003). BETA2/NeuroD1 null mice: a new model for transcription factor-dependent photoreceptor degeneration. J Neurosci 23, 453–461. 44. Yan, R. T., Wang, S. Z. (1998). NeuroD induces photoreceptor cell overproduction in vivo and de novo generation in vitro. J Neurobiol 36, 485–496. 45. Akagi, T., Akita, J., Haruta, M., Suzuki, T., Honda, Y., Inoue, T., Yoshiura, S., Kageyama, R., Yatsu, T., Yamada, M., Takahashi, M. (2005). Iris-derived cells from adult rodents and primates adopt photoreceptor-specific phenotypes. Invest Ophthalmol Vis Sci 46, 3411–3419. 46. Wang, J. C., Harris, W. A. (2005). The role of combinational coding by homeodomain and bHLH transcription factors in retinal cell fate specification. Dev Biol 285, 101–115. 47. Ochocinska, M. J., Hitchcock, P. F. (2007). Dynamic expression of the basic helix-loop-helix transcription factor neuroD in the rod and cone photoreceptor lineages in the retina of the embryonic and larval zebrafish. J Comp Neurol 501, 1–12. 48. Fischer, A. J., Wang, S. Z., Reh, T. A. (2004). NeuroD induces the expression of visinin and calretinin by proliferating cells derived from toxin-damaged chicken retina. Dev Dyn 229, 555–563.

Update on Regulation of Rod Photoreceptor Development

53

49. Hitchcock, P., Kakuk-Atkins, L. (2004). The basic helix-loop-helix transcription factor neuroD is expressed in the rod lineage of the teleost retina. J Comp Neurol 477, 108–117. 50. Yan, R. T., Wang, S. Z. (2004). Requirement of neuroD for photoreceptor formation in the chick retina. Invest Ophthalmol Vis Sci 45, 48–58. 51. Akagi, T., Inoue, T., Miyoshi, G., Bessho, Y., Takahashi, M., Lee, J. E., Guillemot, F., Kageyama, R. (2004). Requirement of multiple basic helix-loop-helix genes for retinal neuronal subtype specification. J Biol Chem 279, 28492–28498. 52. Fossat, N., Chatelain, G., Brun, G., Lamonerie, T. (2006). Temporal and spatial delineation of mouse Otx2 functions by conditional self-knockout. EMBO Rep 7, 824–830. 53. Matsuo, I., Kuratani, S., Kimura, C., Takeda, N., Aizawa, S. (1995). Mouse Otx2 functions in the formation and patterning of rostral head. Genes Dev 9, 2646–2658. 54. Tian, E., Kimura, C., Takeda, N., Aizawa, S., Matsuo, I. (2002). Otx2 is required to respond to signals from anterior neural ridge for forebrain specification. Dev Biol 242, 204–223. 55. Bovolenta, P., Mallamaci, A., Briata, P., Corte, G., Boncinelli, E. (1997). Implication of OTX2 in pigment epithelium determination and neural retina differentiation. J Neurosci 17, 4243–4252. 56. Martinez-Morales, J. R., Dolez, V., Rodrigo, I., Zaccarini, R., Leconte, L., Bovolenta, P., Saule, S. (2003).OTX2activatesthemolecularnetworkunderlyingretinapigmentepitheliumdifferentiation. J Biol Chem 278, 21721–21731. 57. Martinez-Morales, J. R., Signore, M., Acampora, D., Simeone, A., Bovolenta, P. (2001). Otx genes are required for tissue specification in the developing eye. Development 128, 2019–2030. 58. Plouhinec, J. L., Leconte, L., Sauka-Spengler, T., Bovolenta, P., Mazan, S., Saule, S. (2005). Comparative analysis of gnathostome Otx gene expression patterns in the developing eye: implications for the functional evolution of the multigene family. Dev Biol 278, 560–575. 59. Shen, Y. C., Raymond, P. A. (2004). Zebrafish cone-rod (crx) homeobox gene promotes retinogenesis. Dev Biol 269, 237–251. 60. Viczian, A. S., Vignali, R., Zuber, M. E., Barsacchi, G., Harris, W. A. (2003). XOtx5b and XOtx2 regulate photoreceptor and bipolar fates in the Xenopus retina. Development 130, 1281–1294. 61. Nishida, A., Furukawa, A., Koike, C., Tano, Y., Aizawa, S., Matsuo, I., Furukawa, T. (2003). Otx2 homeobox gene controls retinal photoreceptor cell fate and pineal gland development. Nat Neurosci 6, 1255–1263. 62. Baas, D., Bumsted, K. M., Martinez, J. A., Vaccarino, F. M., Wikler, K. C., Barnstable, C. J. (2000). The subcellular localization of Otx2 is cell-type specific and developmentally regulated in the mouse retina. Brain Res Mol Brain Res 78, 26–37. 63. Freund, C. L., Gregory-Evans, C. Y., Furukawa, T., Papaioannou, M., Looser, J., Ploder, L., Bellingham, J., Ng, D., Herbrick, J. A., Duncan, A., Scherer, S. W., Tsui, L. C., Loutradis-Anagnostou, A., Jacobson, S. G., Cepko, C. L., Bhattacharya, S. S., McInnes, R. R. (1997). Cone-rod dystrophy due to mutations in a novel photoreceptor-specific homeobox gene (CRX) essential for maintenance of the photoreceptor. Cell 91, 543–553. 64. Swain, P. K., Chen, S., Wang, Q. L., Affatigato, L. M., Coats, C. L., Brady, K. D., Fishman, G. A., Jacobson, S. G., Swaroop, A., Stone, E., Sieving, P. A., Zack, D. J. (1997). Mutations in the cone-rod homeobox gene are associated with the cone-rod dystrophy photoreceptor degeneration. Neuron 19, 1329–1336. 65. Freund, C. L., Wang, Q. L., Chen, S., Muskat, B. L., Wiles, C. D., Sheffield, V. C., Jacobson, S. G., McInnes, R. R., Zack, D. J., Stone, E. M. (1998). De novo mutations in the CRX homeobox gene associated with Leber congenital amaurosis. Nat Genet 18, 311–312. 66. Jacobson, S. G., Cideciyan, A. V., Huang, Y., Hanna, D. B., Freund, C. L., Affatigato, L. M., Carr, R. E., Zack, D. J., Stone, E. M., McInnes, R. R. (1998). Retinal degenerations with

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69.

70.

71.

72.

73.

74.

75.

76.

77.

78.

79.

80.

81.

Levine and Fuhrmann truncation mutations in the cone-rod homeobox (CRX) gene. Invest Ophthalmol Vis Sci 39, 2417–2426. Perrault, I., Hanein, S., Gerber, S., Barbet, F., Dufier, J. L., Munnich, A., Rozet, J. M., Kaplan, J. (2003). Evidence of autosomal dominant Leber congenital amaurosis (LCA) underlain by a CRX heterozygous null allele. J Med Genet 40, e90. Rivolta, C., Peck, N. E., Fulton, A. B., Fishman, G. A., Berson, E. L., Dryja, T. P. (2001). Novel frameshift mutations in CRX associated with Leber congenital amaurosis. Hum Mutat 18, 550–551. Silva, E., Yang, J. M., Li, Y., Dharmaraj, S., Sundin, O. H., Maumenee, I. H. (2000). A CRX null mutation is associated with both Leber congenital amaurosis and a normal ocular phenotype. Invest Ophthalmol Vis Sci 41, 2076–2079. Sohocki, M. M., Sullivan, L. S., Mintz-Hittner, H. A., Birch, D., Heckenlively, J. R., Freund, C. L., McInnes, R. R., Daiger, S. P. (1998). A range of clinical phenotypes associated with mutations in CRX, a photoreceptor transcription-factor gene. Am J Hum Genet 63, 1307–1315. Swaroop, A., Wang, Q. L., Wu, W., Cook, J., Coats, C., Xu, S., Chen, S., Zack, D. J., Sieving, P. A. (1999). Leber congenital amaurosis caused by a homozygous mutation (R90W) in the homeodomain of the retinal transcription factor CRX: direct evidence for the involvement of CRX in the development of photoreceptor function. Hum Mol Genet 8, 299–305. Furukawa, T., Morrow, E. M., Cepko, C. L. (1997). Crx, a novel otx-like homeobox gene, shows photoreceptor-specific expression and regulates photoreceptor differentiation. Cell 91, 531–541. Chen, S., Wang, Q. L., Nie, Z., Sun, H., Lennon, G., Copeland, N. G., Gilbert, D. J., Jenkins, N. A., Zack, D. J. (1997). Crx, a novel Otx-like paired-homeodomain protein, binds to and transactivates photoreceptor cell-specific genes. Neuron 19, 1017–1030. Bibb, L. C., Holt, J. K., Tarttelin, E. E., Hodges, M. D., Gregory-Evans, K., Rutherford, A., Lucas, R. J., Sowden, J. C., Gregory-Evans, C. Y. (2001). Temporal and spatial expression patterns of the CRX transcription factor and its downstream targets. Critical differences during human and mouse eye development. Hum Mol Genet 10, 1571–1579. Liu, Y., Shen, Y., Rest, J. S., Raymond, P. A., Zack, D. J. (2001). Isolation and characterization of a zebrafish homologue of the cone rod homeobox gene. Invest Ophthalmol Vis Sci 42, 481–487. Sauka-Spengler, T., Baratte, B., Shi, L., Mazan, S. (2001). Structure and expression of an Otx5-related gene in the dogfish Scyliorhinus canicula: evidence for a conserved role of Otx5 and Crxgenes in the specification of photoreceptors. Dev Genes Evol 211, 533–544. Furukawa, A., Koike, C., Lippincott, P., Cepko, C. L., Furukawa, T. (2002). The mouse Crx 5′-upstream transgene sequence directs cell-specific and developmentally regulated expression in retinal photoreceptor cells. J Neurosci 22, 1640–1647. Wu, Y., Cutting, G. R. (2001). Developmentally regulated expression of GABA receptor rho1 and rho2 subunits, L7 and cone-rod homeobox (CRX) genes in mouse retina. Brain Res 912, 1–8. Haruta, M., Kosaka, M., Kanegae, Y., Saito, I., Inoue, T., Kageyama, R., Nishida, A., Honda, Y., Takahashi, M. (2001). Induction of photoreceptor-specific phenotypes in adult mammalian iris tissue. Nat Neurosci 4, 1163–1164. Blackshaw, S., Fraioli, R. E., Furukawa, T., Cepko, C. L. (2001). Comprehensive analysis of photoreceptor gene expression and the identification of candidate retinal disease genes. Cell 107, 579–589. Furukawa, T., Morrow, E. M., Li, T., Davis, F. C., Cepko, C. L. (1999). Retinopathy and attenuated circadian entrainment in Crx-deficient mice. Nat Genet 23, 466–470.

Update on Regulation of Rod Photoreceptor Development

55

82. Livesey, F. J., Furukawa, T., Steffen, M. A., Church, G. M., Cepko, C. L. (2000). Microarray analysis of the transcriptional network controlled by the photoreceptor homeobox gene Crx. Curr Biol 10, 301–310. 83. Morrow, E. M., Furukawa, T., Raviola, E., Cepko, C. L. (2005). Synaptogenesis and outer segment formation are perturbed in the neural retina of Crx mutant mice. BMC Neurosci 6, 5. 84. Chau, K. Y., Chen, S., Zack, D. J., Ono, S. J. (2000). Functional domains of the cone-rod homeobox (CRX) transcription factor. J Biol Chem 275, 37264–37270. 85. Kimura, A., Singh, D., Wawrousek, E. F., Kikuchi, M., Nakamura, M., Shinohara, T. (2000). Both PCE-1/RX and OTX/CRX interactions are necessary for photoreceptor-specific gene expression. J Biol Chem 275, 1152–1160. 86. Lerner, L. E., Peng, G. H., Gribanova, Y. E., Chen, S., Farber, D. B. (2005). Sp4 is expressed in retinal neurons, activates transcription of photoreceptor-specific genes, and synergizes with Crx. J Biol Chem 280, 20642–20650. 87. Mali, R. S., Zhang, X., Hoerauf, W., Doyle, D., Devitt, J., Loffreda-Wren, J., Mitton, K. P. (2007). FIZ1 is expressed during photoreceptor maturation, and synergizes with NRL and CRX at rod-specific promoters in vitro. Exp Eye Res 84, 349–360. 88. Mitton, K. P., Swain, P. K., Chen, S., Xu, S., Zack, D. J., Swaroop, A. (2000). The leucine zipper of NRL interacts with the CRX homeodomain. A possible mechanism of transcriptional synergy in rhodopsin regulation. J Biol Chem 275, 29794–29799. 89. Peng, G. H., Chen, S. (2005). Chromatin immunoprecipitation identifies photoreceptor transcription factor targets in mouse models of retinal degeneration: new findings and challenges. Vis Neurosci 22, 575–586. 90. Qian, J., Esumi, N., Chen, Y., Wang, Q., Chowers, I., Zack, D. J. (2005). Identification of regulatory targets of tissue-specific transcription factors: application to retina-specific gene regulation. Nucleic Acids Res 33, 3479–3491. 91. Yanagi, Y., Takezawa, S., Kato, S. (2002). Distinct functions of photoreceptor cell-specific nuclear receptor, thyroid hormone receptor beta2 and CRX in one photoreceptor development. Invest Ophthalmol Vis Sci 43, 3489–3494. 92. Chau, K. Y., Munshi, N., Keane-Myers, A., Cheung-Chau, K. W., Tai, A. K., Manfioletti, G., Dorey, C. K., Thanos, D., Zack, D. J., Ono, S. J. (2000). The architectural transcription factor high mobility group I(Y) participates in photoreceptor-specific gene expression. J Neurosci 20, 7317–7324. 93. Whitaker, S. L., Knox, B. E. (2004). Conserved transcriptional activators of the Xenopus rhodopsin gene. J Biol Chem 279, 49010–49018. 94. Yanagi, Y., Masuhiro, Y., Mori, M., Yanagisawa, J., Kato, S. (2000). p300/CBP acts as a coactivator of the cone-rod homeobox transcription factor. Biochem Biophys Res Commun 269, 410–414. 95. Otteson, D. C., Liu, Y., Lai, H., Wang, C., Gray, S., Jain, M. K., Zack, D. J. (2004). Kruppel-like factor 15, a zinc-finger transcriptional regulator, represses the rhodopsin and interphotoreceptor retinoid-binding protein promoters. Invest Ophthalmol Vis Sci 45, 2522–2530. 96. Arranz, V., Dreuillet, C., Crisanti, P., Tillit, J., Kress, M., Ernoult-Lange, M. (2001). The zinc finger transcription factor, MOK2, negatively modulates expression of the interphotoreceptor retinoid-binding protein gene, IRBP. J Biol Chem 276, 11963–11969. 97. Otteson, D. C., Lai, H., Liu, Y., Zack, D. J. (2005). Zinc-finger domains of the transcriptional repressor KLF15 bind multiple sites in rhodopsin and IRBP promoters including the CRS-1 and G-rich repressor elements. BMC Mol Biol 6, 15. 98. Wang, X., Xu, S., Rivolta, C., Li, L. Y., Peng, G. H., Swain, P. K., Sung, C. H., Swaroop, A., Berson, E. L., Dryja, T. P., Chen, S. (2002). Barrier to autointegration factor interacts

56

99. 100.

101.

102.

103.

104.

105.

106.

107. 108.

109.

110. 111. 112. 113.

Levine and Fuhrmann with the cone-rod homeobox and represses its transactivation function. J Biol Chem 277, 43288–43300. Zhu, X., Craft, C. M. (2000). Modulation of CRX transactivation activity by phosducin isoforms. Mol Cell Biol 20, 5216–26. Abou-Sleymane, G., Chalmel, F., Helmlinger, D., Lardenois, A., Thibault, C., Weber, C., Merienne, K., Mandel, J. L., Poch, O., Devys, D., Trottier, Y. (2006). Polyglutamine expansion causes neurodegeneration by altering the neuronal differentiation program. Hum Mol Genet 15, 691–703. Chen, S., Peng, G. H., Wang, X., Smith, A. C., Grote, S. K., Sopher, B. L., La Spada, A. R. (2004). Interference of Crx-dependent transcription by ataxin-7 involves interaction between the glutamine regions and requires the ataxin-7 carboxy-terminal region for nuclear localization. Hum Mol Genet 13, 53–67. La Spada, A. R., Fu, Y. H., Sopher, B. L., Libby, R. T., Wang, X., Li, L. Y., Einum, D. D., Huang, J., Possin, D. E., Smith, A. C., Martinez, R. A., Koszdin, K. L., Treuting, P. M., Ware, C. B., Hurley, J. B., Ptacek, L. J., Chen, S. (2001). Polyglutamine-expanded ataxin7 antagonizes CRX function and induces cone-rod dystrophy in a mouse model of SCA7. Neuron 31, 913–927. Palhan, V. B., Chen, S., Peng, G. H., Tjernberg, A., Gamper, A. M., Fan, Y., Chait, B. T., La Spada, A. R., Roeder, R. G. (2005). Polyglutamine-expanded ataxin-7 inhibits STAGA histone acetyltransferase activity to produce retinal degeneration. Proc Natl Acad Sci U S A 102, 8472–8477. Plouhinec, J. L., Sauka-Spengler, T., Germot, A., Le Mentec, C., Cabana, T., Harrison, G., Pieau, C., Sire, J. Y., Veron, G., Mazan, S. (2003). The mammalian Crx genes are highly divergent representatives of the Otx5 gene family, a gnathostome orthology class of orthodenticle-related homeogenes involved in the differentiation of retinal photoreceptors and circadian entrainment. Mol Biol Evol 20, 513–521. Gamse, J. T., Shen, Y. C., Thisse, C., Thisse, B., Raymond, P. A., Halpern, M. E., Liang, J. O. (2002). Otx5 regulates genes that show circadian expression in the zebrafish pineal complex. Nat Genet 30, 117–121. Wang, Q. L., Chen, S., Esumi, N., Swain, P. K., Haines, H. S., Peng, G., Melia, B. M., McIntosh, I., Heckenlively, J. R., Jacobson, S. G., Stone, E. M., Swaroop, A., Zack, D. J. (2004). QRX, a novel homeobox gene, modulates photoreceptor gene expression. Hum Mol Genet 13, 1025–1040. Chen, C. M., Cepko, C. L. (2002). The chicken RaxL gene plays a role in the initiation of photoreceptor differentiation. Development 129, 5363–5375. Ohuchi, H., Tomonari, S., Itoh, H., Mikawa, T., Noji, S. (1999). Identification of chick rax/ rx genes with overlapping patterns of expression during early eye and brain development. Mech Dev 85, 193–195. Pan, Y., Nekkalapudi, S., Kelly, L. E., El-Hodiri, H. M. (2006). The Rx-like homeobox gene (Rx-L) is necessary for normal photoreceptor development. Invest Ophthalmol Vis Sci 47, 4245–4253. Reza, H. M., Yasuda, K. (2004). Roles of Maf family proteins in lens development. Dev Dyn 229, 440–448. Graw, J. (2004). Congenital hereditary cataracts. Int J Dev Biol 48, 1031–1044. Reza, H. M., Urano, A., Shimada, N., Yasuda, K. (2007). Sequential and combinatorial roles of maf family genes define proper lens development. Mol Vis 13, 18–30. Ring, B. Z., Cordes, S. P., Overbeek, P. A., Barsh, G. S. (2000). Regulation of mouse lens fiber cell development and differentiation by the Maf gene. Development 127, 307–317.

Update on Regulation of Rod Photoreceptor Development

57

114. Jamieson, R. V., Perveen, R., Kerr, B., Carette, M., Yardley, J., Heon, E., Wirth, M. G., van Heyningen, V., Donnai, D., Munier, F., Black, G. C. (2002). Domain disruption and mutation of the bZIP transcription factor, MAF, associated with cataract, ocular anterior segment dysgenesis and coloboma. Hum Mol Genet 11, 33–42. 115. Daniele, L. L., Lillo, C., Lyubarsky, A. L., Nikonov, S. S., Philp, N., Mears, A. J., Swaroop, A., Williams, D. S., Pugh, E. N., Jr. (2005). Cone-like morphological, molecular, and electrophysiological features of the photoreceptors of the Nrl knockout mouse. Invest Ophthalmol Vis Sci 46, 2156–2167. 116. Mears, A. J., Kondo, M., Swain, P. K., Takada, Y., Bush, R. A., Saunders, T. L., Sieving, P. A., Swaroop, A. (2001). Nrl is required for rod photoreceptor development. Nat Genet 29, 447–452. 117. Strettoi, E., Mears, A. J., Swaroop, A. (2004). Recruitment of the rod pathway by cones in the absence of rods. J Neurosci 24, 7576–7582. 118. He, L., Campbell, M. L., Srivastava, D., Blocker, Y. S., Harris, J. R., Swaroop, A., Fox, D. A. (1998). Spatial and temporal expression of AP-1 responsive rod photoreceptor genes and bZIP transcription factors during development of the rat retina. Mol Vis 4, 32. 119. Liu, Q., Ji, X., Breitman, M. L., Hitchcock, P. F., Swaroop, A. (1996). Expression of the bZIP transcription factor gene Nrl in the developing nervous system. Oncogene 12, 207–211. 120. O’Brien, K. M., Schulte, D., Hendrickson, A. E. (2003). Expression of photoreceptor-associated molecules during human fetal eye development. Mol Vis 9, 401–409. 121. Swain, P. K., Hicks, D., Mears, A. J., Apel, I. J., Smith, J. E., John, S. K., Hendrickson, A., Milam, A. H., Swaroop, A. (2001). Multiple phosphorylated isoforms of NRL are expressed in rod photoreceptors. J Biol Chem 276, 36824–36830. 122. Akimoto, M., Cheng, H., Zhu, D., Brzezinski, J. A., Khanna, R., Filippova, E., Oh, E. C., Jing, Y., Linares, J. L., Brooks, M., Zareparsi, S., Mears, A. J., Hero, A., Glaser, T., Swaroop, A. (2006). Targeting of GFP to newborn rods by Nrl promoter and temporal expression profiling of flow-sorted photoreceptors. Proc Natl Acad Sci U S A 103, 3890–3895. 123. Oh, E. C., Khan, N., Novelli, E., Khanna, H., Strettoi, E., Swaroop, A. (2007). From the Cover: Transformation of cone precursors to functional rod photoreceptors by bZIP transcription factor NRL. Proc Natl Acad Sci U S A 104, 1679–1684. 124. Bessant, D. A., Holder, G. E., Fitzke, F. W., Payne, A. M., Bhattacharya, S. S., Bird, A. C. (2003). Phenotype of retinitis pigmentosa associated with the Ser50Thr mutation in the NRL gene. Arch Ophthalmol 121, 793–802. 125. Bessant, D. A., Payne, A. M., Mitton, K. P., Wang, Q. L., Swain, P. K., Plant, C., Bird, A. C., Zack, D. J., Swaroop, A., Bhattacharya, S. S. (1999). A mutation in NRL is associated with autosomal dominant retinitis pigmentosa. Nat Genet 21, 355–356. 126. Bessant, D. A., Payne, A. M., Plant, C., Bird, A. C., Swaroop, A., Bhattacharya, S. S. (2000). NRL S50T mutation and the importance of “founder effects” in inherited retinal dystrophies. Eur J Hum Genet 8, 783–787. 127. Martinez-Gimeno, M., Maseras, M., Baiget, M., Beneito, M., Antinolo, G., Ayuso, C., Carballo, M. (2001). Mutations P51L and G122E in retinal transcription factor NRL associated with autosomal dominant and sporadic retinitis pigmentosa. Hum Mutat 17, 520. 128. Nishiguchi, K. M., Friedman, J. S., Sandberg, M. A., Swaroop, A., Berson, E. L., Dryja, T. P. (2004). Recessive NRL mutations in patients with clumped pigmentary retinal degeneration and relative preservation of blue cone function. Proc Natl Acad Sci U S A 101, 17819–17824.

58

Levine and Fuhrmann

129. Robinson-Rechavi, M., Escriva Garcia, H., Laudet, V. (2003). The nuclear receptor superfamily. J Cell Sci 116, 585–586. 130. Ng, L., Hurley, J. B., Dierks, B., Srinivas, M., Salto, C., Vennstrom, B., Reh, T. A., Forrest, D. (2001). A thyroid hormone receptor that is required for the development of green cone photoreceptors. Nat Genet 27, 94–98. 131. Roberts, M. R., Hendrickson, A., McGuire, C. R., Reh, T. A. (2005). Retinoid X receptor (gamma) is necessary to establish the S-opsin gradient in cone photoreceptors of the developing mouse retina. Invest Ophthalmol Vis Sci 46, 2897–2904. 132. Roberts, M. R., Srinivas, M., Forrest, D., Morreale de Escobar, G., Reh, T. A. (2006). Making the gradient: thyroid hormone regulates cone opsin expression in the developing mouse retina. Proc Natl Acad Sci U S A 103, 6218–6223. 133. Srinivas, M., Ng, L., Liu, H., Jia, L., Forrest, D. (2006). Activation of the blue opsin gene in cone photoreceptor development by retinoid-related orphan receptor beta. Mol Endocrinol 20, 1728–1741. 134. Veldhoen, K., Allison, W. T., Veldhoen, N., Anholt, B. R., Helbing, C. C., Hawryshyn, C. W. (2006). Spatio-temporal characterization of retinal opsin gene expression during thyroid hormone-induced and natural development of rainbow trout. Vis Neurosci 23, 169–179. 135. Jacobson, S. G., Marmor, M. F., Kemp, C. M., Knighton, R. W. (1990). SWS (blue) cone hypersensitivity in a newly identified retinal degeneration. Invest Ophthalmol Vis Sci 31, 827–838. 136. Jacobson, S. G., Roman, A. J., Roman, M. I., Gass, J. D., Parker, J. A. (1991). Relatively enhanced S cone function in the Goldmann-Favre syndrome. Am J Ophthalmol 111, 446–453. 137. Marmor, M. F., Jacobson, S. G., Foerster, M. H., Kellner, U., Weleber, R. G. (1990). Diagnostic clinical findings of a new syndrome with night blindness, maculopathy, and enhanced S cone sensitivity. Am J Ophthalmol 110, 124–134. 138. Greenstein, V. C., Zaidi, Q., Hood, D. C., Spehar, B., Cideciyan, A. V., Jacobson, S. G. (1996). The enhanced S cone syndrome: an analysis of receptoral and post-receptoral changes. Vision Res 36, 3711–3722. 139. Hood, D. C., Cideciyan, A. V., Roman, A. J., Jacobson, S. G. (1995). Enhanced S cone syndrome: evidence for an abnormally large number of S cones. Vision Res 35, 1473– 1481. 140. Marmor, M. F., Tan, F., Sutter, E. E., Bearse, M. A., Jr. (1999). Topography of cone electrophysiology in the enhanced S cone syndrome. Invest Ophthalmol Vis Sci 40, 1866–1873. 141. Haider, N. B., Jacobson, S. G., Cideciyan, A. V., Swiderski, R., Streb, L. M., Searby, C., Beck, G., Hockey, R., Hanna, D. B., Gorman, S., Duhl, D., Carmi, R., Bennett, J., Weleber, R. G., Fishman, G. A., Wright, A. F., Stone, E. M., Sheffield, V. C. (2000). Mutation of a nuclear receptor gene, NR2E3, causes enhanced S cone syndrome, a disorder of retinal cell fate. Nat Genet 24, 127–131. 142. Jacobson, S. G., Sumaroka, A., Aleman, T. S., Cideciyan, A. V., Schwartz, S. B., Roman, A. J., McInnes, R. R., Sheffield, V. C., Stone, E. M., Swaroop, A., Wright, A. F. (2004). Nuclear receptor NR2E3 gene mutations distort human retinal laminar architecture and cause an unusual degeneration. Hum Mol Genet 13, 1893–1902. 143. Milam, A. H., Rose, L., Cideciyan, A. V., Barakat, M. R., Tang, W. X., Gupta, N., Aleman, T. S., Wright, A. F., Stone, E. M., Sheffield, V. C., Jacobson, S. G. (2002). The nuclear receptor NR2E3 plays a role in human retinal photoreceptor differentiation and degeneration. Proc Natl Acad Sci U S A 99, 473–478. 144. Sharon, D., Sandberg, M. A., Caruso, R. C., Berson, E. L., Dryja, T. P. (2003). Shared mutations in NR2E3 in enhanced S-cone syndrome, Goldmann-Favre syndrome, and many cases of clumped pigmentary retinal degeneration. Arch Ophthalmol 121, 1316–1323.

Update on Regulation of Rod Photoreceptor Development

59

145. Wright, A. F., Reddick, A. C., Schwartz, S. B., Ferguson, J. S., Aleman, T. S., Kellner, U., Jurklies, B., Schuster, A., Zrenner, E., Wissinger, B., Lennon, A., Shu, X., Cideciyan, A. V., Stone, E. M., Jacobson, S. G., Swaroop, A. (2004). Mutation analysis of NR2E3 and NRL genes in enhanced S cone syndrome. Hum Mutat 24, 439. 146. Bumsted O’Brien, K. M., Cheng, H., Jiang, Y., Schulte, D., Swaroop, A., Hendrickson, A. E. (2004). Expression of photoreceptor-specific nuclear receptor NR2E3 in rod photoreceptors of fetal human retina. Invest Ophthalmol Vis Sci 45, 2807–2812. 147. Akhmedov, N. B., Piriev, N. I., Chang, B., Rapoport, A. L., Hawes, N. L., Nishina, P. M., Nusinowitz, S., Heckenlively, J. R., Roderick, T. H., Kozak, C. A., Danciger, M., Davisson, M. T., Farber, D. B. (2000). A deletion in a photoreceptor-specific nuclear receptor mRNA causes retinal degeneration in the rd7 mouse. Proc Natl Acad Sci U S A 97, 5551–5556. 148. Chen, J., Rattner, A., Nathans, J. (2006). Effects of L1 retrotransposon insertion on transcript processing, localization and accumulation: lessons from the retinal degeneration 7 mouse and implications for the genomic ecology of L1 elements. Hum Mol Genet 15, 2146–2156. 149. Haider, N. B., Demarco, P., Nystuen, A. M., Huang, X., Smith, R. S., McCall, M. A., Naggert, J. K., Nishina, P. M. (2006). The transcription factor Nr2e3 functions in retinal progenitors to suppress cone cell generation. Vis Neurosci 23, 917–929. 150. Takezawa, S., Yokoyama, A., Okada, M., Fujiki, R., Iriyama, A., Yanagi, Y., Ito, H., Takada, I., Kishimoto, M., Miyajima, A., Takeyama, K., Umesono, K., Kitagawa, H., Kato, S. (2007). A cell cycle-dependent co-repressor mediates photoreceptor cell-specific nuclear receptor function. EMBO J 26, 764–774. 151. Turner, D. L., Snyder, E. Y., Cepko, C. L. (1990). Lineage-independent determination of cell type in the embryonic mouse retina. Neuron 4, 833–845. 152. Chen, J., Rattner, A., Nathans, J. (2005). The rod photoreceptor-specific nuclear receptor Nr2e3 represses transcription of multiple cone-specific genes. J Neurosci 25, 118–129. 153. Peng, G. H., Ahmad, O., Ahmad, F., Liu, J., Chen, S. (2005). The photoreceptor-specific nuclear receptor Nr2e3 interacts with Crx and exerts opposing effects on the transcription of rod versus cone genes. Hum Mol Genet 14, 747–764. 154. Corbo, J. C., Cepko, C. L. (2005). A hybrid photoreceptor expressing both rod and cone genes in a mouse model of enhanced S-cone syndrome. PLoS Genet 1, e11. 155. Cheng, H., Khanna, H., Oh, E. C., Hicks, D., Mitton, K. P., Swaroop, A. (2004). Photoreceptor-specific nuclear receptor NR2E3 functions as a transcriptional activator in rod photoreceptors. Hum Mol Genet 13, 1563–75. 156. Cheng, H., Aleman, T. S., Cideciyan, A. V., Khanna, R., Jacobson, S. G., Swaroop, A. (2006). In vivo function of the orphan nuclear receptor NR2E3 in establishing photoreceptor identity during mammalian retinal development. Hum Mol Genet 15, 2588–2602. 157. Haider, N. B., Naggert, J. K., Nishina, P. M. (2001). Excess cone cell proliferation due to lack of a functional NR2E3 causes retinal dysplasia and degeneration in rd7/rd7 mice. Hum Mol Genet 10, 1619–1626. 158. Azadi, S., Zhang, Y., Caffe, A. R., Holmqvist, B., van Veen, T. (2002). Thyroid-beta2 and the retinoid RAR-alpha, RXR-gamma and ROR-beta2 receptor mRNAs; expression profiles in mouse retina, retinal explants and neocortex. Neuroreport 13, 745–750. 159. Hoover, F., Glover, J. C. (1998). Regional pattern of retinoid X receptor-alpha gene expression in the central nervous system of the chicken embryo and its up-regulation by exposure to 9-cis retinoic acid. J Comp Neurol 398, 575–586. 160. Marsh-Armstrong, N., McCaffery, P., Gilbert, W., Dowling, J. E., Drager, U. C. (1994). Retinoic acid is necessary for development of the ventral retina in zebrafish. Proc Natl Acad Sci U S A 91, 7286–7290.

60

Levine and Fuhrmann

161. Mey, J., McCaffery, P., Drager, U. C. (1997). Retinoic acid synthesis in the developing chick retina. J Neurosci 17, 7441–7449. 162. Mori, M., Ghyselinck, N. B., Chambon, P., Mark, M. (2001). Systematic immunolocalization of retinoid receptors in developing and adult mouse eyes. Invest Ophthalmol Vis Sci 42, 1312–1318. 163. Dowling, J. E. (1964). Nutritional and inherited blindness in the rat. Exp Eye Res 15, 348–356. 164. Kelley, M. W., Turner, J. K., Reh, T. A. (1994). Retinoic acid promotes differentiation of photoreceptors in vitro. Development 120, 2091–2102. 165. Stenkamp, D. L., Gregory, J. K., Adler, R. (1993). Retinoid effects in purified cultures of chick embryo retina neurons and photoreceptors. Invest Ophthalmol Vis Sci 34, 2425–2436. 166. Wallace, V. A., Jensen, A. M. (1999). IBMX, taurine and 9-cis retinoic acid all act to accelerate rhodopsin expression in postmitotic cells. Exp Eye Res 69, 617–627. 167. Hyatt, G. A., Schmitt, E. A., Fadool, J. M., Dowling, J. E. (1996). Retinoic acid alters photoreceptor development in vivo. Proc Natl Acad Sci U S A 93, 13298–13303. 168. Kelley, M. W., Williams, R. C., Turner, J. K., Creech-Kraft, J. M., Reh, T. A. (1999). Retinoic acid promotes rod photoreceptor differentiation in rat retina in vivo. Neuroreport 10, 2389–2394. 169. Politi, L. E., Insua, F., Buzzi, E. (1998). Selective outgrowth and differential tropism of amacrine and photoreceptor axons to cell targets during early development in vitro. J Neurosci Res 52, 105–117. 170. Prabhudesai, S. N., Cameron, D. A., Stenkamp, D. L. (2005). Targeted effects of retinoic acid signaling upon photoreceptor development in zebrafish. Dev Biol 287, 157–167. 171. Khanna, H., Akimoto, M., Siffroi-Fernandez, S., Friedman, J. S., Hicks, D., Swaroop, A. (2006). Retinoic acid regulates the expression of photoreceptor transcription factor NRL. J Biol Chem 281, 27327–27334. 172. Li, A., Zhu, X., Brown, B., Craft, C. M. (2003). Gene expression networks underlying retinoic acid-induced differentiation of human retinoblastoma cells. Invest Ophthalmol Vis Sci 44, 996–1007. 173. Soderpalm, A. K., Fox, D. A., Karlsson, J. O., van Veen, T. (2000). Retinoic acid produces rod photoreceptor selective apoptosis in developing mammalian retina. Invest Ophthalmol Vis Sci 41, 937–47. 174. Bernard, M., Klein, D. C. (1996). Retinoic acid increases hydroxyindole-O-methyltransferase activity and mRNA in human Y-79 retinoblastoma cells. J Neurochem 67, 1032–1038. 175. Boatright, J. H., Stodulkova, E., Do, V. T., Padove, S. A., Nguyen, H. T., Borst, D. E., Nickerson, J. M. (2002). The effect of retinoids and butyrate on the expression of CRX and IRBP in retinoblastoma cells. Vision Res 42, 933–938. 176. Li, A., Zhu, X., Craft, C. M. (2002). Retinoic acid upregulates cone arrestin expression in retinoblastoma cells through a cis element in the distal promoter region. Invest Ophthalmol Vis Sci 43, 1375–1383. 177. Liu, H., Mohamed, O., Dufort, D., Wallace, V. A. (2003). Characterization of Wnt signaling components and activation of the Wnt canonical pathway in the murine retina. Dev Dyn 227, 323–334. 178. Van Raay, T. J., Vetter, M. L. (2004). Wnt/frizzled signaling during vertebrate retinal development. Dev Neurosci 26, 352–358. 179. Hsieh, J. C., Kodjabachian, L., Rebbert, M. L., Rattner, A., Smallwood, P. M., Samos, C. H., Nusse, R., Dawid, I. B., Nathans, J. (1999). A new secreted protein that binds to Wnt proteins and inhibits their activities. Nature 398, 431–436.

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180. Hunter, D. D., Zhang, M., Ferguson, J. W., Koch, M., Brunken, W. J. (2004). The extracellular matrix component WIF-1 is expressed during, and can modulate, retinal development. Mol Cell Neurosci 27, 477–488. 181. Liu, H., Thurig, S., Mohamed, O., Dufort, D., Wallace, V. A. (2006). Mapping canonical Wnt signaling in the developing and adult retina. Invest Ophthalmol Vis Sci 47, 5088–5097. 182. Wang, Y., Huso, D., Cahill, H., Ryugo, D., Nathans, J. (2001). Progressive cerebellar, auditory, and esophageal dysfunction caused by targeted disruption of the frizzled-4 gene. J Neurosci 21, 4761–4771. 183. Fu, X., Sun, H., Klein, W. H., Mu, X. (2006). Beta-catenin is essential for lamination but not neurogenesis in mouse retinal development. Dev Biol 299, 424–437. 184. Young, T. L., Cepko, C. L. (2004). A role for ligand-gated ion channels in rod photoreceptor development. Neuron 41, 867–879. 185. Altshuler, D., Lo Turco, J. J., Rush, J., Cepko, C. (1993). Taurine promotes the differentiation of a vertebrate retinal cell type in vitro. Development 119, 1317–1328. 186. Young-Pearse, T. L., Ivic, L., Kriegstein, A. R., Cepko, C. L. (2006). Characterization of mice with targeted deletion of glycine receptor alpha 2. Mol Cell Biol 26, 5728–5734. 187. Bonni, A., Frank, D. A., Schindler, C., Greenberg, M. E. (1993). Characterization of a pathway for ciliary neurotrophic factor signaling to the nucleus. Science 262, 1575–1579. 188. Boulton, T. G., Stahl, N., Yancopoulos, G. D. (1994). Ciliary neurotrophic factor/leukemia inhibitory factor/interleukin 6/oncostatin M family of cytokines induces tyrosine phosphorylation of a common set of proteins overlapping those induced by other cytokines and growth factors. J Biol Chem 269, 11648–11655. 189. Heinrich, P. C., Behrmann, I., Haan, S., Hermanns, H. M., Muller-Newen, G., Schaper, F. (2003). Principles of interleukin (IL)-6-type cytokine signalling and its regulation. Biochem J 374, 1–20. 190. Oh, H., Fujio, Y., Kunisada, K., Hirota, H., Matsui, H., Kishimoto, T., Yamauchi-Takihara, K. (1998). Activation of phosphatidylinositol 3-kinase through glycoprotein 130 induces protein kinase B and p70 S6 kinase phosphorylation in cardiac myocytes. J Biol Chem 273, 9703–9710. 191. Stahl, N., Boulton, T. G., Farruggella, T., Ip, N. Y., Davis, S., Witthuhn, B. A., Quelle, F. W., Silvennoinen, O., Barbieri, G., Pellegrini, S., et al. (1994). Association and activation of Jak-Tyk kinases by CNTF-LIF-OSM-IL-6 beta receptor components. Science 263, 92–95. 192. Fuhrmann, S., Kirsch, M., Hofmann, H. D. (1995). Ciliary neurotrophic factor promotes chick photoreceptor development in vitro. Development 121, 2695–2706. 193. Hicks, D., Molday, R. S. (1986). Differential immunogold-dextran labeling of bovine and frog rod and cone cells using monoclonal antibodies against bovine rhodopsin. Exp Eye Res 42, 55–71. 194. Kirsch, M., Fuhrmann, S., Wiese, A., Hofmann, H. D. (1996). CNTF exerts opposite effects on in vitro development of rat and chick photoreceptors. Neuroreport 7, 697–700. 195. Bradford, R. L., Wang, C., Zack, D. J., Adler, R. (2005). Roles of cell-intrinsic and microenvironmental factors in photoreceptor cell differentiation. Dev Biol 286, 31–45. 196. Xie, H. Q., Adler, R. (2000). Green cone opsin and rhodopsin regulation by CNTF and staurosporine in cultured chick photoreceptors. Invest Ophthalmol Vis Sci 41, 4317–4323. 197. Fuhrmann, S., Heller, S., Rohrer, H., Hofmann, H. D. (1998). A transient role for ciliary neurotrophic factor in chick photoreceptor development. J Neurobiol 37, 672–683. 198. Fuhrmann, S., Kirsch, M., Heller, S., Rohrer, H., Hofmann, H. D. (1998). Differential regulation of ciliary neurotrophic factor receptor-alpha expression in all major neuronal cell classes during development of the chick retina. J Comp Neurol 400, 244–254.

62

Levine and Fuhrmann

199. Hofmann, H. D. (1988). Ciliary neuronotrophic factor stimulates choline acetyltransferase activity in cultured chicken retina neurons. J Neurochem 51, 109–113. 200. Caffe, A. R., Soderpalm, A. K., Holmqvist, I., van Veen, T. (2001). A combination of CNTF and BDNF rescues rd photoreceptors but changes rod differentiation in the presence of RPE in retinal explants. Invest Ophthalmol Vis Sci 42, 275–282. 201. Cayouette, M., Gravel, C. (1997). Adenovirus-mediated gene transfer of ciliary neurotrophic factor can prevent photoreceptor degeneration in the retinal degeneration (rd) mouse. Hum Gene Ther 8, 423–430. 202. LaVail, M. M., Unoki, K., Yasumura, D., Matthes, M. T., Yancopoulos, G. D., Steinberg, R. H. (1992). Multiple growth factors, cytokines, and neurotrophins rescue photoreceptors from the damaging effects of constant light. Proc Natl Acad Sci U S A 89, 11249–11253. 203. Peterson, W. M., Wang, Q., Tzekova, R., Wiegand, S. J. (2000). Ciliary neurotrophic factor and stress stimuli activate the Jak-STAT pathway in retinal neurons and glia. J Neurosci 20, 4081–4090. 204. Sieving, P. A., Caruso, R. C., Tao, W., Coleman, H. R., Thompson, D. J., Fullmer, K. R., Bush, R. A. (2006). Ciliary neurotrophic factor (CNTF) for human retinal degeneration: phase I trial of CNTF delivered by encapsulated cell intraocular implants. Proc Natl Acad Sci U S A 103, 3896–3901. 205. Tao, W., Wen, R., Goddard, M. B., Sherman, S. D., O’Rourke, P. J., Stabila, P. F., Bell, W. J., Dean, B. J., Kauper, K. A., Budz, V. A., Tsiaras, W. G., Acland, G. M., Pearce-Kelling, S., Laties, A. M., Aguirre, G. D. (2002). Encapsulated cell-based delivery of CNTF reduces photoreceptor degeneration in animal models of retinitis pigmentosa. Invest Ophthalmol Vis Sci 43, 3292–3298. 206. Wahlin, K. J., Campochiaro, P. A., Zack, D. J., Adler, R. (2000). Neurotrophic factors cause activation of intracellular signaling pathways in Muller cells and other cells of the inner retina, but not photoreceptors. Invest Ophthalmol Vis Sci 41, 927–936. 207. Wen, R., Song, Y., Kjellstrom, S., Tanikawa, A., Liu, Y., Li, Y., Zhao, L., Bush, R. A., Laties, A. M., Sieving, P. A. (2006). Regulation of rod phototransduction machinery by ciliary neurotrophic factor. J Neurosci 26, 13523–13530. 208. Derouet, D., Rousseau, F., Alfonsi, F., Froger, J., Hermann, J., Barbier, F., Perret, D., Diveu, C., Guillet, C., Preisser, L., Dumont, A., Barbado, M., Morel, A., deLapeyriere, O., Gascan, H., Chevalier, S. (2004). Neuropoietin, a new IL-6-related cytokine signaling through the ciliary neurotrophic factor receptor. Proc Natl Acad Sci U S A 101, 4827–4832. 209. Elliott, J., Cayouette, M., Gravel, C. (2006). The CNTF/LIF signaling pathway regulates developmental programmed cell death and differentiation of rod precursor cells in the mouse retina in vivo. Dev Biol 300, 583–598. 210. Kirsch, M., Hofmann, H. D. (1994). Expression of ciliary neurotrophic factor receptor mRNA and protein in the early postnatal and adult rat nervous system. Neurosci Lett 180, 163–6. 211. Roger, J., Brajeul, V., Thomasseau, S., Hienola, A., Sahel, J. A., Guillonneau, X., Goureau, O. (2006). Involvement of Pleiotrophin in CNTF-mediated differentiation of the late retinal progenitor cells. Dev Biol 298, 527–539. 212. Beltran, W. A., Rohrer, H., Aguirre, G. D. (2005). Immunolocalization of ciliary neurotrophic factor receptor alpha (CNTFRalpha) in mammalian photoreceptor cells. Mol Vis 11, 232–244. 213. Beltran, W. A., Zhang, Q., Kijas, J. W., Gu, D., Rohrer, H., Jordan, J. A., Aguirre, G. D. (2003). Cloning, mapping, and retinal expression of the canine ciliary neurotrophic factor receptor alpha (CNTFRalpha). Invest Ophthalmol Vis Sci 44, 3642–3649.

Update on Regulation of Rod Photoreceptor Development

63

214. Kirsch, M., Lee, M. Y., Meyer, V., Wiese, A., Hofmann, H. D. (1997). Evidence for multiple, local functions of ciliary neurotrophic factor (CNTF) in retinal development: expression of CNTF and its receptors and in vitro effects on target cells. J Neurochem 68, 979–990. 215. Rhee, K. D., Yang, X. J. (2003). Expression of cytokine signal transduction components in the postnatal mouse retina. Mol Vis 9, 715–722. 216. Valter, K., Bisti, S., Stone, J. (2003). Location of CNTFRalpha on outer segments: evidence of the site of action of CNTF in rat retina. Brain Res 985, 169–75. 217. Ozawa, Y., Nakao, K., Shimazaki, T., Takeda, J., Akira, S., Ishihara, K., Hirano, T., Oguchi, Y., Okano, H. (2004). Downregulation of STAT3 activation is required for presumptive rod photoreceptor cells to differentiate in the postnatal retina. Mol Cell Neurosci 26, 258–270. 218. Rhee, K. D., Goureau, O., Chen, S., Yang, X. J. (2004). Cytokine-induced activation of signal transducer and activator of transcription in photoreceptor precursors regulates rod differentiation in the developing mouse retina. J Neurosci 24, 9779–9788. 219. Zhang, S. S., Wei, J., Qin, H., Zhang, L., Xie, B., Hui, P., Deisseroth, A., Barnstable, C. J., Fu, X. Y. (2004). STAT3-mediated signaling in the determination of rod photoreceptor cell fate in mouse retina. Invest Ophthalmol Vis Sci 45, 2407–2412. 220. Ezzeddine, Z. D., Yang, X., DeChiara, T., Yancopoulos, G., Cepko, C. L. (1997). Postmitotic cells fated to become rod photoreceptors can be respecified by CNTF treatment of the retina. Development 124, 1055–1067. 221. Kirsch, M., Schulz-Key, S., Wiese, A., Fuhrmann, S., Hofmann, H. (1998). Ciliary neurotrophic factor blocks rod photoreceptor differentiation from postmitotic precursor cells in vitro. Cell Tissue Res 291, 207–216. 222. Neophytou, C., Vernallis, A. B., Smith, A., Raff, M. C. (1997). Muller-cell-derived leukaemia inhibitory factor arrests rod photoreceptor differentiation at a postmitotic pre-rod stage of development. Development 124, 2345–2354. 223. Schulz-Key, S., Hofmann, H. D., Beisenherz-Huss, C., Barbisch, C., Kirsch, M. (2002). Ciliary neurotrophic factor as a transient negative regulator of rod development in rat retina. Invest Ophthalmol Vis Sci 43, 3099–3108. 224. Cepko, C. L., Austin, C. P., Yang, X., Alexiades, M., Ezzeddine, D. (1996). Cell fate determination in the vertebrate retina. Proc Natl Acad Sci U S A 93, 589–595. 225. Bhattacharya, S., Dooley, C., Soto, F., Madson, J., Das, A. V., Ahmad, I. (2004). Involvement of Ath3 in CNTF-mediated differentiation of the late retinal progenitors. Mol Cell Neurosci 27, 32–43. 226. Zahir, T., Klassen, H., Young, M. J. (2005). Effects of ciliary neurotrophic factor on differentiation of late retinal progenitor cells. Stem Cells 23, 424–432. 227. Graham, D. R., Overbeek, P. A., Ash, J. D. (2005). Leukemia inhibitory factor blocks expression of Crx and Nrl transcription factors to inhibit photoreceptor differentiation. Invest Ophthalmol Vis Sci 46, 2601–2610. 228. Ozawa, Y., Nakao, K., Shimazaki, T., Shimmura, S., Kurihara, T., Ishida, S., Yoshimura, A., Tsubota, K., Okano, H. (2007). SOCS3 is required to temporally fine-tune photoreceptor cell differentiation. Dev Biol 303, 591–600. 229. Young, R. W. (1984). Cell death during differentiation of the retina in the mouse. J Comp Neurol 229, 362–373. 230. Reese, B. E. (2004). Developmental plasticity of photoreceptors. Prog Brain Res 144, 3–19. 231. MacLaren, R. E., Pearson, R. A., MacNeil, A., Douglas, R. H., Salt, T. E., Akimoto, M., Swaroop, A., Sowden, J. C., Ali, R. R. (2006). Retinal repair by transplantation of photoreceptor precursors. Nature 444, 203–207.

64

Levine and Fuhrmann

232. Banin, E., Obolensky, A., Idelson, M., Hemo, I., Reinhardtz, E., Pikarsky, E., Ben-Hur, T., Reubinoff, B. (2006). Retinal incorporation and differentiation of neural precursors derived from human embryonic stem cells. Stem Cells 24, 246–257. 233. Das, A. M., Zhao, X., Ahmad, I. (2005). Stem cell therapy for retinal degeneration: retinal neurons from heterologous sources. Semin Ophthalmol 20, 3–10. 234. Haruta, M. (2005). Embryonic stem cells: potential source for ocular repair. Semin Ophthalmol 20, 17–23. 235. Ikeda, H., Osakada, F., Watanabe, K., Mizuseki, K., Haraguchi, T., Miyoshi, H., Kamiya, D., Honda, Y., Sasai, N., Yoshimura, N., Takahashi, M., Sasai, Y. (2005). Generation of Rx+/Pax6+ neural retinal precursors from embryonic stem cells. Proc Natl Acad Sci U S A 102, 11331–11336. 236. Klassen, H., Kiilgaard, J. F., Zahir, T., Ziaeian, B., Kirov, I., Scherfig, E., Warfvinge, K., Young, M. J. (2007). Progenitor cells from the porcine neural retina express photoreceptor markers after transplantation to the subretinal space of allorecipients. Stem Cells 25, 1222–1230. 237. Lamba, D. A., Karl, M. O., Ware, C. B., Reh, T. A. (2006). Efficient generation of retinal progenitor cells from human embryonic stem cells. Proc Natl Acad Sci U S A 103, 12769– 12774. 238. Liang, L., Yan, R. T., Ma, W., Zhang, H., Wang, S. Z. (2006). Exploring RPE as a source of photoreceptors: differentiation and integration of transdifferentiating cells grafted into embryonic chick eyes. Invest Ophthalmol Vis Sci 47, 5066–5074. 239. Merhi-Soussi, F., Angenieux, B., Canola, K., Kostic, C., Tekaya, M., Hornfeld, D., Arsenijevic, Y. (2006). High yield of cells committed to the photoreceptor fate from expanded mouse retinal stem cells. Stem Cells 24, 2060–2070. 240. Meyer, J. S., Katz, M. L., Kirk, M. D. (2005). Stem cells for retinal degenerative disorders. Ann N Y Acad Sci 1049, 135–145. 241. Sugie, Y., Yoshikawa, M., Ouji, Y., Saito, K., Moriya, K., Ishizaka, S., Matsuura, T., Maruoka, S., Nawa, Y., Hara, Y. (2005). Photoreceptor cells from mouse ES cells by coculture with chick embryonic retina. Biochem Biophys Res Commun 332, 241–247. 242. Sun, G., Asami, M., Ohta, H., Kosaka, J., Kosaka, M. (2006). Retinal stem/progenitor properties of iris pigment epithelial cells. Dev Biol 289, 243–252. 243. Kowalczyk, A. P., Moses, K. (2002). Photoreceptor cells in flies and mammals: Crumby homology? Dev Cell 2, 253–254. 244. Tsujikawa, M., Malicki, J. (2004). Genetics of photoreceptor development and function in zebrafish. Int J Dev Biol 48, 925–934.

4 Photoreceptor–RPE Interactions Physiology and Molecular Mechanisms Silvia C. Finnemann and Yongen Chang CONTENTS Introduction Retinal Adhesion Photoreceptor Outer Segment Renewal Perspective References

INTRODUCTION Photoreceptor rod and cone neurons are highly specialized to detect photons and initiate phototransduction. The machinery for these functions is localized to the outer segment, which faces the apical microvilli-rich surface of the retinal pigment epithelium (RPE) in the subretinal space. It is the function of the RPE to permanently and continuously support photoreceptor function and health through interactions with outer segments. These interactions may be direct, via receptor-ligand pairs that reside on the photoreceptor outer segment (POS) and RPE plasma membranes and link the two, or indirect, via extracellular compounds in the interphotoreceptor matrix (IPM) that act as soluble bridge molecules that are recognized by receptors on both neighboring cell types. Both RPE and photoreceptor cells are normally postmitotic in the healthy adult mammalian eye. Thus, interactions between specific POSs and RPE cells persist for life. Importantly, as RPE cells cover much larger areas of the retina than individual photoreceptor cells, each RPE cell faces between 30 and 50 POSs. Depending on its location in the retina, an individual RPE cell faces and functionally interacts with cones, rods, or (in most cases) a mixture of both. Like the photoreceptors they support, RPE cells are highly polarized cells with distinct protein composition at their apical and basolateral surfaces that face the retina and the choroid connective tissue, respectively. Tight junctions of the RPE serve as permeability barriers that separate apical and basolateral plasma membrane domains to prevent loss of membrane protein and lipid asymmetry by lateral diffusion in the plane of the membrane. From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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Fig. 1. Retinal pigment epithelium (RPE) and photoreceptor contacts in the subretinal space. The subretinal space is separated from the vascularized inner retina and choroidal tissue by tight junctions (t. j.). Tight junctions between photoreceptor inner segments and Müller cells form the external limiting membrane (ELM). Tight junctions of RPE cells separate apical and basolateral domains of the RPE and form the outer blood retinal barrier. Cadherin-based adherens junctions (a. j.) between RPE cells further stabilize the RPE monolayer. Basally, RPE cells adhere to Bruch’s membrane via cell–substrate interactions. The subretinal space contains the interphotoreceptor matrix (IPM) with distinct composition around cones and rods (X and ▲ as indicated), photoreceptor inner segments (IS) and outer segments (OS), and RPE apical membrane with microvilli that extend to ensheathe the photoreceptor OS.

In addition, these multiprotein complexes seal off the subretinal space from the underlying connective tissue to prevent free exchange of extracellular molecules by transepithelial diffusion. Together with the external limiting membrane (ELM) generated by photoreceptorMüller cell junctions, the tight junctions of the RPE form the outer blood-retinal barrier isolating the avascular area of the subretinal space, which contains the apical plasma membrane domain of the RPE, the IPM, and photo-receptor rod and cone outer segments (Fig. 1). Regardless of size and charge, molecules trafficking between the subretinal space and the underlying choroidal circulation must cross the RPE via paracellular or transcellular routes that are largely controlled by RPE cells themselves.

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Concentrations of soluble compounds such as ions, metabolites, and macromolecules in the subretinal space are highly dynamic, varying both spatially and temporally. Changes that are best understood occur as a direct consequence of photoisomerization and phototransduction (recently reviewed in [1]): (1) All-trans retinal released from rhodopsin is reduced to all-trans retinol in the outer segment, which moves through the subretinal space to the RPE for reisomerization. Trafficking of 11-cis retinal from the RPE to the outer segment completes the visual cycle of rod photopigment regeneration. Several specialized retinoid-binding proteins sequester retinoids en route. Indeed, interphotoreceptor retinoid-binding protein (IRBP) is one of the most abundant proteins in the subretinal space [2]. (2) Rhodopsin isomerization in POSs closes cyclic guanosine monophosphate (cGMP)-gated cation channels in the plasma membrane of the outer segment, reducing the dark current that involves flux of Na+ into and K+ out of the outer segment. Thus, photoreceptor illumination directly decreases K+ (from ~5 to 2 mM) and increases Na+ in the subretinal space. These changes in ion concentrations are rapid and transient as ion transporters and channels of the apical surface of RPE cells and of photoreceptor inner segments respond to these changes with activity changes that compensate for them. The distribution and mobility of macromolecules of the IPM in the subretinal space are only poorly understood. Both RPE and photoreceptor cells contribute components to the IPM, generating a regionalized network of numerous glycoproteins and proteoglycans that sequesters small metabolites such as fatty acids and other nutrients [3–7]. Distinct sheaths consisting of high molecular weight proteoglycans and glycoproteins surround cones and rods [8, 9]. These matrix sheaths likely provide important mechanical support for outer segments and RPE apical microvilli that stabilizes their alignment. Moreover, they may serve to bind and sequester nutrients, growth factors, and other proteins for optimal access by either rods or cones [10]. There is much evidence that macromolecule distribution in the subretinal space is tightly controlled. Uehara in the LaVail group first demonstrated in 1990 that the binding pattern of wheat germ agglutinin lectin (WGA) to adult rat retina cross sections depends on illumination. WGA (as well as colloidal iron and antibodies to chondroitin-6-sulfate or IRBP) exhibited staining that was diffusely distributed and restricted to apical and basal zones of the outer segment layer in dark- and light-adapted retina, respectively [11]. The same investigators later showed that recognition sites for peanut agglutinin, which specifically labels components of the cone matrix sheath, do not redistribute in response to differences in illumination [12]. These are very intriguing observations, suggesting that the organization of extracellular macromolecules in the subretinal space may change to facilitate functions specifically associated with rod outer segments, such as the classical visual cycle that employs reisomerization within the RPE. However, the precise molecular nature of these changes and their physiological purpose remain unknown. Taken together, photoreceptor and RPE cells interact in the highly dynamic environment of the subretinal space. In the following, we discuss in detail interactions that take place between POSs and the apical surface of the RPE in the context of this complex extracellular milieu during retinal adhesion and during POS renewal.

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RETINAL ADHESION Physiology of Retinal Adhesion Efficient exchange of substrates between POSs, IPM, and RPE requires maximal alignment and interdigitation. The apical surface of the RPE, including microvilli, tightly adheres to IPM components and possibly photoreceptors in the healthy retina, maintaining this tissue organization. The importance of retinal adhesion to retinal function becomes obvious in the drastic consequences of retinal detachment. Persistent retinal detachment directly results in RPE dedifferentiation and proliferation, POS degeneration, and photoreceptor cell death [13]. Furthermore, retinal detachment causes irreversible changes in inner retinal neurons. For a comprehensive recent review of these effects of retinal detachment on retinal cells, please see [14]. In the context of RPE–photoreceptor interactions, it is particularly relevant that IPM proteoglycan rearrangement and RPE microvilli collapse are early, reversible responses to retinal detachment. A variety of physiological activities contribute to retinal adhesion. These include active fluid transport, intraocular pressure, and osmotic pressure gradients. In addition, apical surface receptors of the RPE adhere to specific IPM ligands [15]. The actin-binding protein ezrin is an important and abundant structural component of the apical microvilli of the RPE [16]. Ezrin expression is both necessary and sufficient for RPE microvilli extension [17, 18]. At the light microscopic level, ezrin distribution in the mammalian retina does not change with illumination or time of day [19]. This suggests that apical RPE microvilli extension and interdigitate in between POSs is constant in mammalian retina. In contrast, the strength of retinal adhesion as measured by RPE attachment to peeled-off neural retina (see Molecular Mechanisms of Retinal Adhesion and Fig. 2) increases by 58% between 2 and 3.5 h after light onset in mice entrained to a 12-h light/12-h dark cycle [19]. Retinal adhesion increases on time even in constant darkness in previously entrained mice. This suggests that murine retinal adhesion does not respond to light but may be regulated by circadian rhythms. In contrast, light onset directly increases retinal adhesion in Xenopus laevis retina [20]. Finally, strength of retinal adhesion decreases with age in mouse retina such that resistance of RPE to shear stress in 12-month-old mice is only 45% compared to the resistance in 1-monthold retina in animals of the same genotype [19]. The molecular basis of this change in RPE–photoreceptor interaction remains to be explored. Molecular Mechanisms of Retinal Adhesion Studying the molecular mechanisms of retinal adhesion in the healthy retina is hampered by the lack of experimental model systems. Because of the numerous participating components and their complicated and highly organized architecture, neither direct nor indirect, IPM-mediated adhesive interactions of RPE and POSs have been reconstituted ex vivo. However, Endo and colleagues in 1988 developed and characterized an assay testing strength of RPE–retinal adhesion in enucleated rabbit eyes [21] (Fig. 2). These investigators established that RPE adhesion directly correlates with the amount of RPE pigment fractionating with the neural retina when it is peeled from an open eyecup. Using this assay, Endo and colleagues found that postmortem, retinal adhesion in rabbit

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Fig. 2. Mechanical separation of retinal pigment epithelium (RPE) and neural retina allows quantification of retinal adhesion. In a modification of a protocol by Endo and colleagues [21], mice are sacrificed by CO2 asphyxiation. Lens and cornea are swiftly removed from each enucleated eyeball (A) in HEPES-buffered Hank’s saline solution containing Ca2+ and Mg2+ at room temperature, conditions that preserve retinal adhesion. After transferring an individual eyecup to an empty plastic dish, a single radial cut is performed toward the optic nerve, flattening the eyecup retina facing up. B The neural retina is then peeled off with forceps from one side of the cut to the other. C The isolated retina reveals attached apical portions of RPE cells that contain melanin pigment. D Bright-field microscopy of a whole-mounted peeled-off retina with exposed outer retinal surface demonstrates the extent of RPE retrieval with retina harvested from 2-month-old wild-type mice 2 h after light onset. Scale bar 100 µm (Printed with permission from [19].)

retina rapidly decreases if tissues are chilled on ice or incubated in Ca2+- and Mg2+-free buffer solution. This suggests that receptors involved in retinal adhesion require physiological temperature and the presence of divalent cations for ligand binding. Focusing on the role of glycosylated IPM components on retinal adhesion, Yao and colleagues subsequently found that subretinal injections of chondroitinase ABC, neuraminidase, and testicular hyaluronidase temporarily reduce retinal adhesion, suggesting that regulation of retinal adhesion in vivo may involve changes in glycosylation [22]. A similar experimental approach showed that the drug cytochalasin D weakens

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retinal adhesion in a concentration-dependent manner if applied to the subretinal space, suggesting a role for actin microfilaments in RPE–photoreceptor interactions [23]. While some progress has been made in characterizing the complex mixture of glycoproteins and proteoglycans molecules that comprise the IPM, we know l ittle about RPE surface receptors that participate in retinal adhesion. The homotypic neural-cell adhesion molecule (N-CAM) localizes to the apical surface of RPE and to outer segments, suggesting that it may mediate direct interactions between the two cell types [24]. However, this hypothesis remains to be tested directly. Most recently, the increasing availability of mutant animal models has facilitated study of the contributions of individual molecules to retinal adhesion. For example, vitiligo mice, which carry a mutation in the microphthalmia transcription factor gene [25], display early onset retinal detachment likely due to a primary defect in retinal adhesion [26]. Expression of specific adhesive receptors or ligands that participate in retinal adhesion may thus be reduced in vitiligo mice. However, this possibility has not yet been investigated. The cell–substrate adhesion receptor αvβ5 integrin is the only receptor of the large integrin family that localizes to the apical surface of the RPE [27–29]. The apical surface of the RPE is the sole site of αvβ5 integrin expression in the retina. Localization of αvβ5 integrin at the RPE’s phagocytic surface is conserved between rats, mice, and humans [27, 28]. Comparing strength of retinal adhesion between β5 integrin knockout mice and strain- and age-matched, wild-type mice served to directly assess whether lack of apical αvβ5 integrin altered retinal adhesion. In a modification of the protocol of Endo and colleagues (Fig. 2), retina samples were solubilized in detergent buffer following mechanical peeling off the eyecup. This method of preparation allowed quantifying both RPE pigment and RPE and retinal proteins in each sample by spectroscopy and by immunoblotting, respectively. Importantly, melanin quantification of neural retina samples correlates very closely with partitioning of the RPE-specific cytoplasmic protein RPE65 and the RPE apical microvilli marker protein ezrin with the neural retina, confirming that RPE fractionation with neural retina accurately reflects RPE–retinal adhesion. As shown in Fig. 3, the experiments demonstrated greatly reduced pigment as well as proteins RPE65 and ezrin retrieved with the ripped-off neural retina at all times of day in β5 integrin knockout mice compared to strain- and age-matched wild-type mice. Thus, retinal adhesion is severely weakened in β5 integrin knockout mice. In addition, lack of αvβ5 greatly attenuates the synchronized daily fluctuation of retinal adhesion (Fig. 3B). Thus, αvβ5 integrin receptors contribute to retinal adhesion at all times and have a major role in maximizing retinal adhesion 3.5 h after light onset. β5 integrin knockout retina contain wild-type levels of other receptor proteins of the RPE thought to be relevant for retinal adhesion, N-CAM and integrin subunits other than β5 integrin’s partner subunit αv. This suggests that lack of αvβ5 integrin receptors decreases retinal adhesion directly. These experiments identified αvβ5 integrin as the first RPE surface receptor directly implicated in retinal adhesion. In summary, the recent identification of the role of αvβ5 integrin in retinal adhesion has confirmed the long-standing hypothesis that specific apical plasma membrane receptors of the RPE adhere to IPM components or outer segment surface receptors via cell–substrate or cell–cell interactions, respectively. Given the repertoire of extracellular glycoproteins

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Fig. 3. Decreased strength and attenuated diurnal peak of retinal adhesion in β5 integrin knockout mice. A Bright-field microscopy shows exposed outer retinal surface of peeled-off retinas harvested from 2-month-old mice at 8 a.m. β5 integrin knockout retina retains significantly less retinal pigment epithelium (RPE) pigment compared to wild-type retina. Scale 100 µm B Immunoblots of lysates prepared from individual peeled-off neural retina samples harvested from 2-month-old mice at indicated times of day show increased amounts of the RPE marker protein RPE65 in samples harvested at 9.30 a.m. compared with other time points. Ezrin, MerTK (tyrosine kinase Mer), and IRBP (interphotoreceptor retinol-binding protein) did not change significantly. Notably, RPE65 levels in β5 integrin knockout retina samples also increased between 9:30 a.m. and 8 a.m. but remained far below levels of RPE65 in wild-type retina samples at all times of day. (Modified with permission from [19].)

that serve as ligands of αvβ5 and related integrins in other tissues, it appears likely that there are specific ligands for αvβ5 integrin in the IPM that bridge the RPE–retina interface. These ligands, as well as RPE receptors that presumably contribute to retinal adhesion in addition to αvβ5, remain to be identified. Significance of Retinal Adhesion for Retinal Function Long-term retinal detachment causes outer segment degeneration [30] and, subsequently, apoptotic cell death of photoreceptors [13], proving that proper retinal attachment is critical for vision. Despite their obvious importance for photoreceptor survival and hence vision, we still know little about RPE surface receptors for IPM ligands that mediate

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retinal adhesion. Recent insight into the role of αvβ5 integrin in retinal adhesion gained by exploring RPE pigment and marker protein fractionation and localization in β5 integrin knockout mice has demonstrated an experimental strategy that may prove useful in identifying additional receptor proteins of both RPE and photoreceptor and their subretinal ligands in the near future. Our better understanding of the molecular mechanisms of retinal adhesion are much needed to develop new therapeutic strategies aiming to prevent irreversible vision loss following retinal detachment by providing a microenvironment in the subretinal space that maximally promotes recovery of both damaged photoreceptors and RPE cells. PHOTORECEPTOR OUTER SEGMENT RENEWAL Physiology of Outer Segment Disk Assembly and Disk Shedding Photoreceptor neurons normally do not renew themselves in the adult mammalian retina. However, Bairati and Orzalesi first hypothesized in 1963 that both rods and cones continuously turn over their outer segment portions, replacing the membrane disks that carry rhodopsin/cone opsins and the phototransduction machinery [31]. Since Young in 1967 provided the first direct evidence for such a process, now termed outer segment renewal [32], such de novo formation of outer segment disks has been extensively studied in numerous species [33, 34]. Photoreceptor cells synthesize outer segment resident proteins and lipids mainly in their inner segment. Following translation in the rough endoplasmic reticulum, nascent polypeptides mature while passing through the Golgi apparatus and traffic to the outer segment from the trans-Golgi network packaged in transport vesicles [35, 36]. Vesicles traverse the connecting cilium to contribute to the assembly of new membrane disks by evagination at the proximal end of the outer segment [35, 37]. Cones principally differ from rods with respect to protein turnover: Cone disks do not separate from each other, allowing diffusion of newly synthesized and assembled proteins within the length of the outer segment [38, 39]. In contrast, individual rod disks vary in age, with those in the proximal end of the outer segment most recently generated and those at the distal tip facing the RPE most aged. Outer segment protein biosynthesis follows a daily rhythm that is in part regulated at the transcript level, as amounts of rhodopsin and cone opsin messenger RNAs (mRNAs) are maximal at the onset of dark [40–42]. Circadian rhythms upregulate gene transcription in chick cone photoreceptors to raise iodopsin (red cone opsin) mRNA levels prior to the onset of the dark period [43]. It has not yet been reported whether rates of transcription or transcript stability increase to reach higher steady-state levels of specific mRNAs in mammalian retina. In amphibian retina, outer segment disk assembly does not occur evenly at all times but is regulated by illumination and circadian rhythms [33, 44, 45]. In contrast, illumination has little effect on rates of disk assembly in mouse rods [46]. Photoreceptor disk assembly is generally thought to be largely photoreceptor cell autonomous and independent of photoreceptor–RPE interactions. However, in the healthy retina, production of new POS disks is in a precise balance with elimination of most aged portions of outer segments to maintain constant length of outer segments with time. Young and Bok (1969) first demonstrated the disposal of distal, most aged

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rod outer segment tips in packets of uniform size at the interface of outer segments with the RPE in frog retina, a process that became to be known as disk shedding [47]. Unlike disk assembly, shedding of POS distal fragments most likely requires active participation of directly underlying RPE cells. RPE apical microvilli ensheathe rod and cone outer segments, and their extensions can be observed in between shedding POS and the remainder of the outer segment [48]. Moreover, outer segments cease to shed their distal tips in detached Xenopus laevis retina [49]. Rod POS shedding is highly synchronized in the retina and occurs at light onset in most species, regulated by circadian rhythms in mammals [50, 51], by illumination in frogs (Rana pipiens) [52], and by a combination of both in Xenopus laevis [53]. In the mouse retina, each rod photoreceptor sheds a packet of disks comprising approximately 10% of its total length once a day [46]. Similar rates of rod shedding are documented for other species. Newly synthesized protein distributes diffusely throughout the length of the outer segment of photoreceptor cones [54]. Therefore, distal cone outer segment disks do not contain most aged outer segment components as distal rod disks do. Nevertheless, cones shed distal tips of their outer segments in similar size fragments to rods regardless of species and retinal location [55]. In lizard, goldfish, and chick retina, cone photoreceptors shed after the onset of dark [56–58]. In ground squirrel retina, cones shed either following the onset of dark or in the middle of the dark period [59]. In the cat [60] and in the diurnal Nile rat [61] retina, cones, like rods, shed following the onset of light. Taken together, cones shed distal tips of their outer segments in synchronized bursts like rods, but there is considerable variability among species with regard to the timing of cone shedding. Physiology of RPE Engulfment of Shed Outer Segment Fragments The synchronicity of rod disk aging and shedding facilitated autoradiographic tracer studies that allowed Young and Bok in 1969 to demonstrate that RPE cells phagocytose shed rod POS [47]. Furthermore, the Royal College of Surgeons (RCS) rat strain has long provided an animal model of hereditary retinal degeneration caused by abnormal accumulation of POS in the subretinal space [62]. Mullen and LaVail generated and studied chimeric rats with mosaic RCS and wild-type RPE to demonstrate that the presence of wild-type RPE underlying RCS photoreceptor rods was sufficient to promote normal POS turnover. Therefore, POS accumulation in the RCS retina occurs as a consequence of deficiencies in the RPE rather than in photoreceptors [63]. The rapid and complete degeneration of the retina in the RCS rat illustrates the importance of the RPE and its phagocytic activity in POS renewal specifically and in retinal homeostasis in general. The rhythmic and continuous nature of POS phagocytosis in the retina renders RPE cells the most active phagocytes in nature. The enormous task of the RPE cells in POS disposal becomes immediately obvious when one considers that each RPE cell in the mammalian retina faces numerous photoreceptor rods (e.g., ~45 in the peripheral rhesus monkey retina and a staggering 300 in the rat retina; [64]), each one of which sheds its distal tip containing about 100 disks every morning. Each RPE cell must therefore completely dispose of several thousand outer segment disks before its next phagocytic challenge. Since RPE cells do not turn over in the adult mammalian eye, each individual cell must clear its enormous phagocytic load promptly and efficiently every 24 h over many decades.

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The RPE cells in the healthy mammalian retina respond to circadian rod POS disk shedding with a vigorous phagocytic response and then cease phagocytosis until the next shedding event. Indeed, electron microscopy images of the retina–RPE interface acquired from samples of tissues harvested at times of POS shedding usually reveal POS that still remain attached to the outer segment or POS that were already internalized by the RPE. Observation of shed POS that reside in the subretinal space awaiting RPE engulfment is very rare. This suggests that RPE cells may be optimally prepared at light onset to respond to shed POS with prompt internalization. At other times, intact outer segment distal tips remain in immediate proximity to RPE apical projections without triggering phagocytic attack. Taken together, this suggests that the phagocytic function of the RPE may be tightly temporally regulated in the retina. The rhythmic nature of RPE clearance of POS requires that RPE cells rid themselves of phagocytosed material before the next phagocytic challenge to avoid gradual accumulation of POS with time. Despite its importance in maintaining RPE long-term function, few studies have focused on digestion of POS. Lysosomes of different size and enzymatic content fuse with POS phagosomes by as early as 30 min after light onset [65]. Electron microscopy images show that the disk structure of engulfed POS in RPE phagosomes disappears gradually starting at around 2.5 h after light onset in light-entrained rats [65]. At the light microscopic level, numbers of discernible phagosomes in albino mouse RPE return to baseline levels approximately 3 h after light onset [66]. Finally, at least some of the final products of POS digestion, such as docosahexaenoic acid, are transported back for reuse by photoreceptors [67–71]. Molecular Mechanisms of Shedding and RPE Phagocytosis Studies seeking to identify the molecular machinery used by photoreceptors and RPE cells for POS renewal have greatly benefited from the fact that RPE cells in tissue culture retain their phagocytic activity toward POS. Recording the binding and internalization kinetics of POS by RPE in culture ideally complements the classical microscopic characterization of outer segment uptake by RPE in vivo (Fig. 4). First, outer segment recognition cannot be studied separately from outer segment internalization in vivo because shed outer segments in the subretinal space are juxtaposed to the RPE surface whether or not they are recognized or bound by RPE receptors. Second, far greater numbers of RPE cells can be evaluated in each sample in assays in vitro than in tissue sections. Therefore, small but significant alterations in RPE phagocytic activity may be detected by in vitro assays that may be missed in the light and electron microscopy studies of postmortem tissues. Third, RPE cells in vitro can be studied following specific manipulation of their phagocytic mechanism by pharmacological compounds, recombinant proteins, protein overexpression or downregulation, just to name a few. Gain- and loss-of-function approaches are well suited to unequivocally identify critical components of the RPE phagocytic machinery. Fourth, in vitro phagocytic challenge of RPE cells allows one to test directly the phagocytic activity of RPE cells toward POS, while altered photoreceptor shedding or IPM may indirectly alter the phagocytic activity by RPE cells in vivo. Importantly, however, RPE cells in vitro only provide a phagocytic assay system with relevance to RPE phagocytosis in vivo if cells are studied as differentiated, polarized epithelial monolayers that assemble their phagocytic machinery at their apical surface,

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Fig. 4. Fluorescence microscopy quantification of photoreceptor outer segment (POS) phagocytosis by retinal pigment epithelial (RPE) cells in vivo and in vitro. A–C Cryosections of eyecups from 2-month-old wild-type mice were labeled with rod opsin antibody B6-30 [105]. Maximal projections of confocal microscopy x–y sections representing tissue sections 5-µm thick are shown. A High-magnification view of the RPE–POS interface shows intact rod outer segments and opsin-positive phagosomes (white) in RPE cells adjacent to RPE nuclei (gray). B and C Lowmagnification view of similar opsin signals without nuclei stain illustrates that these images can be used to count opsin phagosome numbers in the RPE. B At 1 h before light onset, the RPE cell layer shows few opsin-labeled phagosomes. C At 2 h after light onset, the RPE cell layer shows numerous opsin-labeled phagosomes, confirming the daily burst of rod POS phagocytosis by the RPE. D Maximal projection of confocal microscopy x–y sections of primary wild-type mouse RPE cells in culture 1 h after phagocytic challenge demonstrates vigorous uptake of fluorescent isolated bovine POS by RPE cells in vitro. Phagocytosed POS appears in white, RPE cell junctions stained with ZO-1 tight junction marker antibody appear in gray. All scale bars 20 µm.

like RPE cells in vivo. Finally, while all evidence suggests that the phagocytic activity of polarized RPE cells in vitro retains the primary characteristics of the phagocytic activity of RPE cells in vivo, this is not the case for the nature of particle contact. In experimental phagocytosis assays, RPE cells must establish firm binding of isolated POS that is stable enough to withstand shear forces during sample processing, including vigorous washing steps. This is in sharp contrast to the contact of apical RPE receptors with shedding/shed POS in the subretinal space, where mechanical stress is absent and a stable binding event per se may not occur. Thus, comparison of in vivo and in vitro RPE phagocytosis counting fluorescence- or opsin antibody-labeled POS as illustrated in Fig. 4 are both required to fully elucidate the phagocytic machinery of the RPE. The events that cause and accompany shedding of photoreceptor tips remain largely obscure. As outlined in the previous section, it is generally thought that RPE microvilli– outer segment interactions as well as photoreceptor intrinsic mechanisms contribute to

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POS shedding. However, the molecular mechanisms that promote outer segment shedding remain yet to be uncovered. This includes the identification of putative “shed me” and “eat me” signals exposed by photoreceptor tips. Molecular changes likely designate the distal tip for shedding and may also serve as recognition signals triggering subsequent RPE phagocytosis. POS changes in surface structure or composition may promote recognition by RPE phagocytic receptors directly. Alternatively, POS changes may promote recruitment of ligands, concentrating or presenting it for engagement of RPE surface receptors. This scenario predicts that the same ligand–ligand complex binds to POS and to RPE receptors. Although there is no evidence for such ligand as yet for retina, a precedent exists. Most cells undergoing apoptosis flip the anionic phospholipid phosphatidylserine from the inner to the outer leaflet of the plasma membrane lipid bilayer [72]. External phosphatidylserine serves to signal the apoptotic process to phagocytic cells. Phagocytes may express surface receptors that recognize phosphatidylserine directly [73]. In addition, soluble phosphatidylserine-binding proteins form a bridge between apoptotic and phagocytic cells. It has not yet been demonstrated whether shedding POS in the retina exposes external leaflet phosphatidylserine. However, phosphatidylserine-enriched liposomes have been reported to affect in vitro outer segment uptake [74]. Furthermore, apoptotic cells and POS compete for binding by RPE cells and by macrophages in culture [75]. This strongly suggests that these different phagocytic particles share “eat me” signals. The only such signal conclusively identified for apoptotic cells is phosphatidylserine. Much progress has been made over the past decade identifying the components that form the RPE phagocytic machinery. Most important, it is now clear that RPE cells use a mechanism to phagocytose shed outer segment that belongs to a group of related clearance mechanisms used by other cell types, such as macrophages, to phagocytose apoptotic cells [75, 76]. Professional phagocytic mechanisms like the RPE’s are saturable, receptormediated processes with distinct phases of particle recognition/binding, internalization, and digestion. All known phagocytic mechanisms employ multiple phagocyte receptors, five of which have been described in RPE cells as described below. In vitro evidence suggests that pattern recognition receptors also associated with inflammatory phagocytosis may participate in POS uptake. Inhibiting a mannose-binding protein using antibodies reduces POS uptake by rat RPE in culture [77]. Toll-like receptor 4 has been shown to redistribute at the apical surface of human RPE cells in response to POS isolated from human but not from bovine retina and to induce a cytoplasmic signaling response in the RPE that may be involved in triggering POS clearance [78]. Little is known yet about the significance of these receptors for POS renewal in vivo. The scavenger receptor family member CD36 facilitates apoptotic cell clearance, fatty acid transport, and cell–matrix interactions by recognizing structurally defined lipid peroxidation products [79–82]. Blocking CD36 with CD36 antibodies partially inhibits outer segment engulfment by RPE cells in vitro [83]. Clustering of CD36 is sufficient to alter the rate with which RPE cells in culture internalize surface-bound outer segment [84]. In vitro assays suggest that CD36 clustering may activate intracellular signaling processes that ultimately target the internalization mechanism of the RPE. Lack of functional or morphological retinal abnormalities in CD36 knockout mice suggests that CD36 may be dispensable for POS renewal in the healthy eye, at least in mice kept under standard vivarium conditions. However, acute high-intensity illumination generates oxidized

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phospholipids in rat retina that are specifically recognized by CD36 [85]. In vitro, these ligands impair POS phagocytosis by RPE cells in a CD36-dependent manner. Notably, rats that harbor a deletion variant in the scavenger receptor CD36 were found to be more sensitive to intense light-induced retinal damage [86]. Thus, CD36 signaling may not be essential for RPE phagocytosis under basal conditions but may be involved in outer segment clearance under conditions that increase oxidative stress in the retina. A deletion in the coding region of the gene for the receptor tyrosine kinase Mer (MerTK) abolishes RPE phagocytosis in the RCS rat [87, 88]. MerTK deficiency also causes rapid retinal degeneration and defective apoptotic cell clearance by macrophages in MerTK kinase-deficient transgenic mice [76, 89]. MerTK-deficient RCS RPEs in culture retain the ability to recognize and bind outer segment but fail to internalize surface-bound outer segments [90, 91]. Despite the critical importance of MerTK for outer segment phagocytosis in vitro and in vivo, its downstream effectors in RPE cells remain to be identified. The permanent RPE cell lines RPE-J (rat), ARPE-19 and d407 (both human), and mouse, rat, and human RPE in primary culture employ the integrin adhesion receptor αvβ5 to recognize and bind isolated POS in experimental phagocytosis assays [28, 92– 94]. The αvβ5 integrin is essential for POS binding in vitro as RPE cells derived from β5 integrin knockout mice in primary culture largely fail to bind isolated POS [95]. RPE cells in β5 integrin knockout mice in vivo retain basal levels of phagocytic activity but lack the characteristic burst of phagocytosis upon early morning rod shedding (Fig. 5A) [95]. Moreover, αvβ5 integrin deficiency is sufficient to cause age-related vision loss in β5 integrin knockout mice accompanied by excessive accumulation of lipofuscin lipid storage bodies in the RPE, a cardinal feature of RPE aging and disease [95]. These findings illustrate that αvβ5 integrin receptors of the RPE are critical for retinal function. Furthermore, studies comparing wild-type RPE cells with RPE cells lacking αvβ5 receptor in vivo and in vitro indicated that αvβ5 integrin receptor engagement initiates intracellular signaling processes in the RPE that promote POS engulfment. Integrin receptors do not possess intrinsic enzymatic activity. However, integrin cytoplasmic domains assemble cytosolic signaling proteins and often bind different sets of proteins depending on receptor occupancy. Indeed, phagocytic challenge with POS rapidly increases the presence of multiple tyrosine-phosphorylated proteins associated with apical αvβ5 integrin in cultured RPE cells [96]. One of these proteins is focal adhesion kinase (FAK) [96]. FAK is a cytoplasmic tyrosine kinase that relays integrin signals [97]. Dramatic reduction of POS internalization by RPE cells in which FAK is specifically inhibited demonstrates that FAK is an important mediator of RPE phagocytic signaling downstream of αvβ5 integrin. MerTK tyrosine phosphorylation is thought to reflect MerTK activity. Strikingly, silencing FAK signaling in RPE cells abolishes MerTK tyrosine phosphorylation induced by POS phagocytosis [96]. MerTK is thus a target of FAK signaling in RPE cells. Furthermore, a strict temporal regulation of both FAK and MerTK activities precisely coincides with circadian shedding of rod outer segments in intact mouse retina [95]. These synchronized signaling events are completely abolished in αvβ5 integrindeficient retina [95] (Fig. 5B). These data provide the first direct evidence of a crosstalk of the recognition receptor αvβ5 integrin and the internalization receptor MerTK: The αvβ5 integrin-dependent signaling via FAK controls the efficiency of RPE phagocytosis by promoting peak MerTK activity at the time of rod outer segment shedding.

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Fig. 5. Loss of the daily burst of retinal pigment epithelium (RPE) phagocytosis and synchronized activation of FAK (focal adhesion kinase) and MerTK (tyrosine kinase Mer) in mice lacking αvβ5 integrin. A Phagosome quantification reveals that β5 integrin knockout mouse retina lacks the characteristic burst of phagocytosis that follows light onset at 6 a.m. in wildtype mouse retina. Bars represent mean numbers of phagosomes ± standard deviation (n = 3). B Immunoblotting compares protein expression and phosphorylation profiles in eyecup detergent lysates harvested at different times of day. Promptly after light onset at 6 a.m., levels of active, phosphorylated FAK and MerTK increase in wild-type but not in β5 integrin knockout mouse eyecups. The rise in FAK phosphorylation precedes the increase of MerTK phosphorylation. This agrees well with earlier data from in vitro phagocytosis assays showing that MerTK activation during photoreceptor outer segment (POS) uptake requires FAK activation [96]. (Modified with permission from [95].)

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Taken together, all available evidence indicates that αvβ5 integrin at the apical surface of the RPE has important functions in POS phagocytosis as well as in retinal adhesion. These functions may be largely independent of each other since both are defective immediately following the establishment of mature RPE–photoreceptor interactions in the β5 knockout mouse retina. It is possible that αvβ5 receptors at the same plasma membrane domain of the RPE exist in two independent functional pools that mediate retinal adhesion and POS phagocytosis, respectively. Notably, αvβ5 integrin-dependent FAK-MerTK signaling coincides with the daily burst of rod POS phagocytosis but not with the daily peak of retinal adhesion [19, 95]. Thus, the two distinct functions of αvβ5 receptors at the apical surface of the RPE may utilize different downstream signaling pathways to promote two distinct aspects of RPE–photoreceptor interactions. Significance of Photoreceptor Outer Segment Renewal for Retinal Function Photoreceptor function strictly depends on efficient RPE phagocytosis of spent OS. Complete failure of RPE cells to engulf POS causes the rapid photoreceptor degeneration in the RCS rat [63, 98, 99]. Furthermore, the continuous nature of outer segment renewal implies that delay in POS removal will gradually cause POS components to accumulate. Indeed, lipofuscin storage bodies containing a complex mix of lipid and protein buildup in human RPE over time as a direct consequence of incomplete POS digestion [100]. Delayed outer segment opsin degradation in transgenic mice causes lipofuscin accumulation and retinal dysfunction with age [101]. Loss of synchronicity of outer segment clearance in β5 integrin null mice is sufficient to promote accumulation of lipofuscin and loss of photoreceptor function with age [95]. Lipofuscin components likely directly decrease RPE function and viability [94, 102]. Finally, there is much evidence that impaired RPE phagocytosis also contributes to human retinal disease, such as retinitis pigmentosa and age-related macular degeneration [103, 104]. PERSPECTIVE Numerous interactions between RPE and photoreceptor cells take place at all times in the retina. Important physiological functions of these interactions, such as retinal adhesion and POS renewal, occur at the same site and sometimes simultaneously. In addition, RPE–photoreceptor interactions are highly dynamic and differ among retinal regions with the circadian rhythm or with illumination. Although important recent progress has been made, many aspects of these interactions remain poorly understood at the molecular level. This is particularly true with regard to specific interactions of the RPE with photoreceptor cones. Given the enormous significance of cone function for human vision, it is thus imperative to elucidate the molecular mechanisms of renewal and adhesion, specifically of cone photoreceptors, in the mammalian retina in future studies. ACKNOWLEDGMENTS We apologize to those colleagues whose work we did not cite due to space restrictions. This work was supported by National Institutes of Health grants EY13295 and EY17173. S.C.F. is the recipient of a William and Mary Greve Special Scholar Award by Research to Prevent Blindness and of an Irma T. Hirschl Career Award.

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REFERENCES 1. Strauss O. The retinal pigment epithelium in visual function. Physiol Rev 2005;85(3):845–881. 2. Pfeffer B, Wiggert B, Lee L, Zonnenberg B, Newsome D, Chader G. The presence of a soluble interphotoreceptor retinol-binding protein (IRBP) in the retinal interphotoreceptor space. J Cell Physiol 1983;117(3):333–341. 3. Johnson LV, Hageman GS. Structural and compositional analyses of isolated cone matrix sheaths. Invest Ophthalmol Vis Sci 1991;32(7):1951–1957. 4. Johnson LV, Hageman GS, Blanks JC. Restricted extracellular matrix domains ensheath cone photoreceptors in vertebrate retinae. Prog Clin Biol Res 1985;190:33–44. 5. Hollyfield JG, Rayborn ME, Landers RA, Myers KM. Insoluble interphotoreceptor matrix domains surround rod photoreceptors in the human retina. Exp Eye Res 1990;51(1):107–110. 6. Hollyfield JG. Hyaluronan and the functional organization of the interphotoreceptor matrix. Invest Ophthalmol Vis Sci 1999;40(12):2767–2769. 7. Hollyfield JG, Varner HH, Rayborn ME. Regional variation within the interphotoreceptor matrix from fovea to the retinal periphery. Eye 1990;4(Pt 2):333–339. 8. Johnson LV, Hageman GS, Blanks JC. Interphotoreceptor matrix domains ensheath vertebrate cone photoreceptor cells. Invest Ophthalmol Vis Sci 1986;27(2):129–135. 9. Hollyfield JG, Varner HH, Rayborn ME, Osterfeld AM. Retinal attachment to the pigment epithelium. Retina 1989;9(1):59–68. 10. Steinberg RH. Interactions between the retinal pigment epithelium and the neural retina. Doc Ophthalmol 1985;60(4):327–346. 11. Uehara F, Matthes MT, Yasumura D, LaVail MM. Light-evoked changes in the interphotoreceptor matrix. Science 1990;248(4963):1633–1636. 12. Uehara F, Yasumura D, LaVail MM. Rod- and cone-associated interphotoreceptor matrix in the rat retina. Differences in light-evoked distributional changes. Invest Ophthalmol Vis Sci 1991;32(2):285–292. 13. Cook B, Lewis GP, Fisher SK, Adler R. Apoptotic photoreceptor degeneration in experimental retinal detachment. Invest Ophthalmol Vis Sci 1995;36(6):990–996. 14. Fisher SK, Lewis GP, Linberg KA, Verardo MR. Cellular remodeling in mammalian retina: results from studies of experimental retinal detachment. Prog Retin Eye Res 2005;24(3):395–431. 15. Hageman GS, Marmor MF, Yao XY, Johnson LV. The interphotoreceptor matrix mediates primate retinal adhesion. Arch Ophthalmol 1995;113(5):655–660. 16. Bonilha VL, Bhattacharya SK, West KA, et al. Support for a proposed retinoid-processing protein complex in apical retinal pigment epithelium. Exp Eye Res 2004;79(3):419–422. 17. Bonilha VL, Rayborn ME, Saotome I, McClatchey AI, Hollyfield JG. Microvilli defects in retinas of ezrin knockout mice. Exp Eye Res 2006;82(4):720–729. 18. Bonilha VL, Finnemann SC, Rodriguez-Boulan E. Ezrin promotes morphogenesis of apical microvilli and basal infoldings in retinal pigment epithelium. J Cell Biol 1999;147(7): 1533–1548. 19. Nandrot EF, Anand M, Sircar M, Finnemann SC. Novel role for αvβ5 integrin in retinal adhesion and its diurnal peak. Am J Physiol Cell Physiol 2006;290(4):C1256–C1262. 20. Defoe DM, Matsumoto B, Besharse JC. Cytochalasin D inhibits L-glutamate-induced disc shedding without altering L-glutamate-induced increase in adhesiveness. Exp Eye Res 1989;48(5):641–652. 21. Endo EG, Yao XY, Marmor MF. Pigment adherence as a measure of retinal adhesion: dependence on temperature. Invest Ophthalmol Vis Sci 1988;29(9):1390–1396. 22. Yao XY, Hageman GS, Marmor MF. Retinal adhesiveness is weakened by enzymatic modification of the interphotoreceptor matrix in vivo. Invest Ophthalmol Vis Sci 1990;31(10):2051–2058.

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23. Chiang RK, Yao XY, Takeuchi A, Dalal R, Marmor MF. Cytochalasin D reversibly weakens retinal adhesiveness. Curr Eye Res 1995;14(12):1109–1113. 24. Gundersen D, Powell SK, Rodriguez-Boulan E. Apical polarization of N-CAM in retinal pigment epithelium is dependent on contact with the neural retina. J Cell Biol 1993;121(2):335–343. 25. Lamoreux ML, Boissy RE, Womack JE, Nordlund JJ. The vit gene maps to the mi (microphthalmia) locus of the laboratory mouse. J Hered 1992;83(6):435–439. 26. Bora N, Defoe D, Smith SB. Evidence of decreased adhesion between the neural retina and retinal pigmented epithelium of the Mitfvit (vitiligo) mutant mouse. Cell Tissue Res 1999;295(1):65–75. 27. Anderson DH, Johnson LV, Hageman GS. Vitronectin receptor expression and distribution at the photoreceptor-retinal pigment epithelial interface. J Comp Neurol 1995;360(1):1–16. 28. Finnemann SC, Bonilha VL, Marmorstein AD, Rodriguez-Boulan E. Phagocytosis of rod outer segments by retinal pigment epithelial cells requires αvβ5 integrin for binding but not for internalization. Proc Natl Acad Sci U S A 1997;94(24):12932–12937. 29. Clegg DO, Mullick LH, Wingerd KL, et al. Adhesive events in retinal development and function: the role of integrin receptors. Results Probl Cell Differ 2000;31:141–156. 30. Machemer R, Norton EW. Experimental retinal detachment in the owl monkey. Am J Ophthalmol 1968;66(3):396–427, 1075–1091. 31. Bairati A Jr, Orzalesi N. The ultrastructure of the pigment epithelium and of the photoreceptor-pigment epithelium junction in the human retina. J Ultrastruct Res 1963;41:484–496. 32. Young RW. The renewal of photoreceptor cell outer segments. J Cell Biol 1967;33(1):61–72. 33. Hollyfield JG, Rayborn ME, Verner GE, Maude MB, Anderson RE. Membrane addition to rod photoreceptor outer segments: light stimulates membrane assembly in the absence of increased membrane biosynthesis. Invest Ophthalmol Vis Sci 1982;22(4):417–427. 34. Papermaster DS, Converse CA, Siuss J. Membrane biosynthesis in the frog retina: opsin transport in the photoreceptor cell. Biochemistry 1975;14(7):1343–1352. 35. Young RW, Droz B. The renewal of protein in retinal rods and cones. J Cell Biol 1968;39(1):169–184. 36. Papermaster DS, Schneider BG, Besharse JC. Vesicular transport of newly synthesized opsin from the Golgi apparatus toward the rod outer segment. Ultrastructural immunocytochemical and autoradiographic evidence in Xenopus retinas. Invest Ophthalmol Vis Sci 1985;26(10):1386–1404. 37. Steinberg RH, Fisher SK, Anderson DH. Disc morphogenesis in vertebrate photoreceptors. J Comp Neurol 1980;190(3):501–508. 38. Bok D, Young RW. The renewal of diffusely distributed protein in the outer segments of rods and cones. Vision Res 1972;12(2):161–168. 39. Poo M, Cone RA. Lateral diffusion of rhodopsin in the photoreceptor membrane. Nature 1974;247(441):438–441. 40. Korenbrot JI, Fernald RD. Circadian rhythm and light regulate opsin mRNA in rod photoreceptors. Nature 1989;337(6206):454–457. 41. Kamphuis W, Cailotto C, Dijk F, Bergen A, Buijs RM. Circadian expression of clock genes and clock-controlled genes in the rat retina. Biochem Biophys Res Commun 2005;330(1):18–26. 42. von Schantz M, Lucas RJ, Foster RG. Circadian oscillation of photopigment transcript levels in the mouse retina. Brain Res Mol Brain Res 1999;72(1):108–114. 43. Pierce ME, Sheshberadaran H, Zhang Z, Fox LE, Applebury ML, Takahashi JS. Circadian regulation of iodopsin gene expression in embryonic photoreceptors in retinal cell culture. Neuron 1993;10(4):579–584. 44. Besharse JC, Hollyfield JG, Rayborn ME. Photoreceptor outer segments: accelerated membrane renewal in rods after exposure to light. Science 1977;196(4289):536–538.

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Finnemann and Chang

45. Matsumoto B, Bok D. Diurnal variations in amino acid incorporation into inner segment opsin. Invest Ophthalmol Vis Sci 1984;25(1):1–9. 46. Besharse JC, Hollyfield JG. Turnover of mouse photoreceptor outer segments in constant light and darkness. Invest Ophthalmol Vis Sci 1979;18(10):1019–1024. 47. Young RW, Bok D. Participation of the retinal pigment epithelium in the rod outer segment renewal process. J Cell Biol 1969;42(2):392–403. 48. Steinberg RH, Wood I, Hogan MJ. Pigment epithelial ensheathment and phagocytosis of extrafoveal cones in human retina. Philos Trans R Soc Lond B Biol Sci 1977;277(958): 459–474. 49. Williams DS, Fisher SK. Prevention of rod disk shedding by detachment from the retinal pigment epithelium. Invest Ophthalmol Vis Sci 1987;28(1):184–187. 50. LaVail MM. Circadian nature of rod outer segment disc shedding in the rat. Invest Ophthalmol Vis Sci 1980;19(4):407–411. 51. LaVail MM. Rod outer segment disk shedding in rat retina: relationship to cyclic lighting. Science 1976;194(4269):1071–1074. 52. Basinger S, Hoffman R, Matthes M. Photoreceptor shedding is initiated by light in the frog retina. Science 1976;194(4269):1074–1076. 53. Besharse JC, Hollyfield JG, Rayborn ME. Turnover of rod photoreceptor outer segments. II. Membrane addition and loss in relationship to light. J Cell Biol 1977;75(2 Pt 1):507–527. 54. Young RW. An hypothesis to account for a basic distinction between rods and cones. Vision Res 1971;11(1):1–5. 55. Anderson DH, Fisher SK, Steinberg RH. Mammalian cones: disc shedding, phagocytosis, and renewal. Invest Ophthalmol Vis Sci 1978;17(2):117–133. 56. O’Day WT, Young RW. Rhythmic daily shedding of outer-segment membranes by visual cells in the goldfish. J Cell Biol 1978;76(3):593–604. 57. Young RW. The daily rhythm of shedding and degradation of cone outer segment membranes in the lizard retina. J Ultrastruct Res 1977;61(2):172–185. 58. Young RW. The daily rhythm of shedding and degradation of rod and cone outer segment membranes in the chick retina. Invest Ophthalmol Vis Sci 1978;17(2):105–116. 59. Long KO, Fisher SK, Fariss RN, Anderson DH. Disc shedding and autophagy in the conedominant ground squirrel retina. Exp Eye Res 1986;43(2):193–205. 60. Fisher SK, Pfeffer BA, Anderson DH. Both rod and cone disc shedding are related to light onset in the cat. Invest Ophthalmol Vis Sci 1983;24(7):844–856. 61. Bobu C, Craft CM, Masson-Pevet M, Hicks D. Photoreceptor organization and rhythmic phagocytosis in the Nile rat Arvicanthis ansorgei: a novel diurnal rodent model for the study of cone pathophysiology. Invest Ophthalmol Vis Sci 2006;47(7):3109–3118. 62. Bourne MC, Campbell DA, Tansley K. Hereditary degeneration of the rat retina. Br J Ophthalmol 1938;22:613–622. 63. Mullen RJ, LaVail MM. Inherited retinal dystrophy: primary defect in pigment epithelium determined with experimental rat chimeras. Science 1976;192(4241):799–801. 64. Besharse J, Defoe D. The role of the retinal pigment epithelium in photoreceptor membrane turnover. In: Marmor MF, Wolfensberger TJ, eds. The retinal pigment epithelium. New York: Oxford University Press; 1998:152–172. 65. Bosch E, Horwitz J, Bok D. Phagocytosis of outer segments by retinal pigment epithelium: phagosome-lysosome interaction. J Histochem Cytochem 1993;41(2):253–263. 66. Gibbs D, Kitamoto J, Williams DS. Abnormal phagocytosis by retinal pigmented epithelium that lacks myosin VIIa, the Usher syndrome 1B protein. Proc Natl Acad Sci U S A 2003;100:6481–6486.

Photoreceptor–RPE Interactions

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67. Gordon WC, Bazan NG. Docosahexaenoic acid utilization during rod photoreceptor cell renewal. J Neurosci 1990;10(7):2190–2202. 68. Rodriguez de Turco EB, Parkins N, Ershov AV, Bazan NG. Selective retinal pigment epithelial cell lipid metabolism and remodeling conserves photoreceptor docosahexaenoic acid following phagocytosis. J Neurosci Res 1999;57(4):479–486. 69. Gordon WC, Rodriguez de Turco EB, Bazan NG. Retinal pigment epithelial cells play a central role in the conservation of docosahexaenoic acid by photoreceptor cells after shedding and phagocytosis. Curr Eye Res 1992;11(1):73–83. 70. Stinson AM, Wiegand RD, Anderson RE. Recycling of docosahexaenoic acid in rat retinas during n-3 fatty acid deficiency. J Lipid Res 1991;32(12):2009–2017. 71. Wiegand RD, Koutz CA, Stinson AM, Anderson RE. Conservation of docosahexaenoic acid in rod outer segments of rat retina during n-3 and n-6 fatty acid deficiency. J Neurochem 1991;57(5):1690–1699. 72. Fadeel B. Plasma membrane alterations during apoptosis: role in corpse clearance. Antioxid Redox Signal 2004;6(2):269–275. 73. Fadok VA, Bratton DL, Rose DM, Pearson A, Ezekewitz RA, Henson PM. A receptor for phosphatidylserine-specific clearance of apoptotic cells. Nature 2000;405(6782):85–90. 74. Ryeom SW, Silverstein RL, Scotto A, Sparrow JR. Binding of anionic phospholipids to retinal pigment epithelium may be mediated by the scavenger receptor CD36. J Biol Chem 1996;271(34):20536–20539. 75. Finnemann SC, Rodriguez-Boulan E. Macrophage and retinal pigment epithelium phagocytosis: apoptotic cells and photoreceptors compete for αvβ3 and αvβ5 integrins, and protein kinase C regulates αvβ5 binding and cytoskeletal linkage. J Exp Med 1999;190(6):861–874. 76. Scott RS, McMahon EJ, Pop SM, et al. Phagocytosis and clearance of apoptotic cells is mediated by MER. Nature 2001;411(6834):207–211. 77. Boyle D, Tien LF, Cooper NG, Shepherd V, McLaughlin BJ. A mannose receptor is involved in retinal phagocytosis. Invest Ophthalmol Vis Sci 1991;32(5):1464–1470. 78. Kindzelskii AL, Elner VM, Elner SG, Yang D, Hughes BA, Petty HR. Toll-like receptor 4 (TLR4) of retinal pigment epithelial cells participates in transmembrane signaling in response to photoreceptor outer segments. J Gen Physiol 2004;124(2):139–149. 79. Ren Y, Silverstein RL, Allen J, Savill J. CD36 gene transfer confers capacity for phagocytosis of cells undergoing apoptosis. J Exp Med 1995;181(5):1857–1862. 80. Jimenez B, Volpert OV, Crawford SE, Febbraio M, Silverstein RL, Bouck N. Signals leading to apoptosis-dependent inhibition of neovascularization by thrombospondin-1. Nat Med 2000;6(1):41–48. 81. Silverstein RL, Febbraio M. CD36 and atherosclerosis. Curr Opin Lipidol 2000;11(5): 483–491. 82. Podrez EA, Poliakov E, Shen Z, et al. A novel family of atherogenic oxidized phospholipids promotes macrophage foam cell formation via the scavenger receptor CD36 and is enriched in atherosclerotic lesions. J Biol Chem 2002;277(41):38517–38523. 83. Ryeom SW, Sparrow JR, Silverstein RL. CD36 participates in the phagocytosis of rod outer segments by retinal pigment epithelium. J Cell Sci 1996;109(Pt 2):387–395. 84. Finnemann SC, Silverstein RL. Differential roles of CD36 and αvβ5 integrin in photoreceptor phagocytosis by the retinal pigment epithelium. J Exp Med 2001;194(9):1289–1298. 85. Sun M, Finnemann SC, Febbraio M, et al. Light-induced oxidation of photoreceptor outer segment phospholipids generates ligands for CD36-mediated phagocytosis by retinal pigment epithelium: a potential mechanism for modulating outer segment phagocytosis under oxidant stress conditions. J Biol Chem 2006;281(7):4222–4230.

86

Finnemann and Chang

86. Li S, Lam TT, Fu J, Tso MO. Systemic hypertension exaggerates retinal photic injury. Arch Ophthalmol 1995;113(4):521–526. 87. D’Cruz PM, Yasumura D, Weir J, et al. Mutation of the receptor tyrosine kinase gene Mertk in the retinal dystrophic RCS rat. Hum Mol Genet 2000;9(4):645–651. 88. Nandrot E, Dufour EM, Provost AC, et al. Homozygous deletion in the coding sequence of the c-mer gene in RCS rats unravels general mechanisms of physiological cell adhesion and apoptosis. Neurobiol Dis 2000;7(6 Pt B):586–599. 89. Duncan JL, LaVail MM, Yasumura D, et al. An RCS-like retinal dystrophy phenotype in mer knockout mice. Invest Ophthalmol Vis Sci 2003;44(2):826–838. 90. Edwards RB, Szamier RB. Defective phagocytosis of isolated rod outer segments by RCS rat retinal pigment epithelium in culture. Science 1977;197(4307):1001–1003. 91. Chaitin MH, Hall MO. Defective ingestion of rod outer segments by cultured dystrophic rat pigment epithelial cells. Invest Ophthalmol Vis Sci 1983;24(7):812–820. 92. Miceli MV, Newsome DA, Tate DJ, Jr. Vitronectin is responsible for serum-stimulated uptake of rod outer segments by cultured retinal pigment epithelial cells. Invest Ophthalmol Vis Sci 1997;38(8):1588–1597. 93. Lin H, Clegg DO. Integrin avb5 participates in the binding of photoreceptor rod outer segments during phagocytosis by cultured human retinal pigment epithelium. Invest Ophthalmol Vis Sci 1998;39(9):1703–1712. 94. Finnemann SC, Leung LW, Rodriguez-Boulan E. The lipofuscin component A2E selectively inhibits phagolysosomal degradation of photoreceptor phospholipid by the retinal pigment epithelium. Proc Natl Acad Sci U S A 2002;99(6):3842–3847. 95. Nandrot EF, Kim Y, Brodie SE, Huang X, Sheppard D, Finnemann SC. Loss of synchronized retinal phagocytosis and age-related blindness in mice lacking αvβ5 integrin. J Exp Med 2004;200:1539–1545. 96. Finnemann SC. Focal adhesion kinase signaling promotes phagocytosis of integrin-bound photoreceptors. EMBO J 2003;22(16):4143–4154. 97. Parsons JT. Focal adhesion kinase: the first ten years. J Cell Sci 2003;116(Pt 8):1409–1416. 98. Dowling JE, Sidman RL. Inherited retinal dystrophy of the rat. J Cell Biol 1962;14:73–109. 99. Bok D, Hall MO. The role of the pigment epithelium in the etiology of inherited retinal dystrophy in the rat. J Cell Biol 1971;49:664–682. 100. Feeney L. Lipofuscin and melanin of human retinal pigment epithelium. Fluorescence, enzyme cytochemical, and ultrastructural studies. Invest Ophthalmol Vis Sci 1978;17(7): 583–600. 101. Rakoczy PE, Zhang D, Robertson T, et al. Progressive age-related changes similar to age-related macular degeneration in a transgenic mouse model. Am J Pathol 2002;161(4): 1515–1524. 102. Sparrow JR, Parish CA, Hashimoto M, Nakanishi K. A2E, a lipofuscin fluorophore, in human retinal pigmented epithelial cells in culture. Invest Ophthalmol Vis Sci 1999;40(12):2988– 2995. 103. Gal A, Li Y, Thompson DA, et al. Mutations in MERTK, the human orthologue of the RCS rat retinal dystrophy gene, cause retinitis pigmentosa. Nat Genet 2000;26(3):270–271. 104. Zareparsi S, Buraczynska M, Branham KE, et al. Toll-like receptor 4 variant D299G is associated with susceptibility to age-related macular degeneration. Hum Mol Genet 2005;14(11):1449–1455. 105. Adamus G, Zam ZS, Arendt A, Palczewski K, McDowell JH, Hargrave PA. Anti-rhodopsin monoclonal antibodies of defined specificity: characterization and application. Vision Res 1991;31(1):17–31.

5 Molecular Biology of IRBP and Its Role in the Visual Cycle Diane E. Borst, Jeffrey H. Boatright, and John M. Nickerson CONTENTS Introduction IRBP Protein Studies IRBP Null Mice IRBP Induces Experimental Autoimmune Uveitis IRBP Expression During Development Variability in IRBP Expression Molecular Biology of IRBP IRBP Genomic Cloning Evolution of IRBP Importance of the STUDY of the Control of Gene Expression Identification of DNA CIS-Acting Controlling Elements: In Vitro and In Vivo Experiments Transcription Factors and Their Role in the Control of IRBP Expression Transgenic Mice Repressors of IRBP Gene Expression Summary and Conjecture References

INTRODUCTION Interphotoreceptor retinoid-binding protein (IRBP; Unigene RBP3) is found in the eye and is critical for vision. It is a 144-kDa glycolipoprotein that is synthesized by the photoreceptor cells and secreted into the interphotoreceptor space (IPS), where it facilitates the transport of retinoids between the retinal pigment epithelium (RPE) and the photoreceptor cells. IRBP, the only retinoid-binding protein found in the IPS, is critical for vision because free retinoids damage cells; yet, retinoids must traverse the IPS twice during the visual cycle. IRBP is a chaperone protein for retinoids. IRBP is known

From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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to induce experimental autoimmune uveitis (EAU) in mice and rats. EAU serves as an animal model for several sight-threatening ocular inflammatory diseases. Increasing our knowledge of the normal expression of IRBP and its regulation will further our understanding of normal retinal cell biology. IRBP PROTEIN STUDIES During the visual cycle, retinoids must traverse the IPM twice. Unbound retinoids can be easily degraded. Also, they are cytotoxic. Retinoid-binding proteins act as chaperones to retinoids, binding them and preventing either of these untoward effects [1]. IRBP is the only retinoid-binding protein known to be present in the IPS and is the major soluble protein found in the IPS [2]. IRBP is a 144-kDa glycolipoprotein that is synthesized by the retina, is present in a number of different vertebrate classes, and is secreted into the IPS. It is known to bind retinoids (e.g., 11-cis retinal, alltrans retinol, and retinoic acid) and fatty acids. Using Western blot analyses on dissected tissue extracts, IRBP is found to be most abundant in interphotoreceptor matrix (IPM) washes, but it is also found in the retina cytosol, aqueous, vitreous, pineal gland cytosol, and the cerebral cortex cytosol [3]. It was not present in the cytosols prepared from the cornea, lens, liver, testes, Harderian gland, or cerebrospinal fluid [3]. Immunoelectron microscopic staining has localized IRBP to the photoreceptor cells and pinealocytes [4]. IRBP NULL MICE IRBP may function in the visual system via its retinoid properties [5–16]. Several investigators have found that IRBP impedes the flow of retinoids from the RPE (or an equivalent vesicle-like source) to the photoreceptors or outer segments. Corresponding in vitro experiments in the transport of retinoids from one type of artificial vesicle to another favor a buffering model of the function of IRBP. Many of these experiments reflect the notion that there are several sinks, pools, or reservoirs of retinoids, and that under different physiological conditions, the connections from one pool to the next is rate limiting. Further experiments are needed to determine the following: (1) Are there additional points where IRBP may participate in regulating retinoid cycling? (2) Is there a receptor for IRBP on either the RPE apical membrane or the plasma membrane of the rod or cone photoreceptor cell? (3) How does IRBP extract retinoids from the RPE or outer segment plasma membrane? IRBP knockout mice were constructed several years ago [17]. Homozygous mice lacking the IRBP gene exhibit a normal life span and have no obvious abnormal phenotypic characteristics outside the eye. Heterozygous mice appear normal as well. The absence of IRBP has no apparent deleterious effect on the rate of retinoid shuttling between the RPE and the photoreceptor cell, an essential process known as the visual cycle [17, 18]. This is not to imply that visual function in IRBP knockout mice is normal. On the contrary, the absence of IRBP yields a visual system with reduced a-wave magnitude, as is demonstrated by electroretinography. This corresponds proportionally to a reduced thickness of the outer nuclear layer (ONL) of roughly 50% as is seen in histological

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sections. IRBP knockout mice appear to have half the a-wave signal and about half the number of rod photoreceptor cells when compared to wild-type mice [17, 18]. The outer segments of the IRBP null mice appear slightly disorganized, and a few small vacuoles are found in the IPS between outer segments [18]. These knockout mouse studies suggest a function for IRBP in the development and maintenance of the photoreceptors. It is not yet clear how or when the absence of IRBP gene expression affects the number of photoreceptor cells. It is not known whether this effect is manifested at the level of the number of rod cells that are born (that is, too few rods being born) or whether too many rod cells die prematurely during retina development. It is fairly obvious by simple inspection of the published images of the IRBP knockout retinas that this defect is not a change in the fate of the cells being born as there is no increase in the number of other kinds of cells in the retina. Similarly, simple inspection also suggests that there is no corresponding diminution in the thicknesses of the other layers of the retina, suggesting that there is no general reduction in the number of bipolar, amacrine, horizontal, Müller glial, or ganglion cells. This decrease in the number of photoreceptor cells but not of the other retinal cells might point to the potential for abnormal development of synapse formation in the knockout mouse, although this has not been reported in the literature. Preliminary findings suggest that the manifestations of the absence of IRBP occur early in the development of the retina as changes are detected at or before P10 by both morphology and microarray analyses [19]. IRBP INDUCES EXPERIMENTAL AUTOIMMUNE UVEITIS Experimental autoimmune uveitis (EAU) serves as an animal model for the study of autoimmune diseases that lead to retinal degeneration. IRBP has been shown to induce EAU in rats, mice, and monkeys when it is injected into the footpad of the animal (for review, see [20]). The pathology of IRBP-induced disease is restricted to the retina and pineal gland: Photoreceptors are missing or degenerated while the rest of the retina remains intact. The disease characteristics are similar to those seen in human uveitis, thus making it an excellent model for study. Specific IRBP peptides are known to be uveitogenic in rats and mice and have been shown to have a common sequence for both the immunological and immunopathological sites [21–26]. IRBP EXPRESSION DURING DEVELOPMENT The neural retina is a well-ordered laminar structure consisting of alternating layers of cells and their synapses. The outermost layers contain the photoreceptor cells. The innermost layer of the retina is the nerve fiber layer containing ganglion cell axons that travel through the optic nerve and carry the visual signals to the brain. Early in development, the retina consists of a single layer of retinoprogenitor cells that, in time, proliferate to form enough cells to comprise all of the different cell types of the retina. A single progenitor cell can generate all of the different retinal cell types [27–30]. During development, populations of progenitor cells stop dividing and differentiate into the different retinal cell types, including both neurons and glia. The different retinal cell types are not all generated at the same time.

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Differentiation occurs in a defined, specific sequence. Developing retinal cells begin to differentiate between embryonic day 10 (E10) and 11 (in mice and rats) into different cell types: ganglion cells, horizontal cells, or cone photoreceptors (mice [31], rats [32]). Cone cells are the first photoreceptors to differentiate in the retina, comprise 3–5% of the photoreceptor population in mouse, and first appear at (E10) (mice [33]; rats [32]). Their differentiation is complete before birth. In mice, rods first appear at E13, and their generation continues until postnatal day 7 (P7) [33]. Rod photoreceptor genesis begins and ends a few days later in rats [32]. In mice, rats, humans, and presumably other mammals, IRBP expression begins early in retinal development. Using Western blot analysis, the presence of the mouse IRBP protein was identified as early as E17, but IRBP was not detected at E15 [34]. Using the more sensitive technique of ribonuclease (RNase) protections analysis, Liou et al. [35] identified low levels of IRBP message in mice at E11.5 and suggested that IRBP may be expressed as early as E10.5. The early expression of IRBP in the mouse retina was confirmed by reverse transcriptase-coupled polymerase chain reaction [rtPCR] experiments that show an amplification product at E10.5 but not E9.5 [36]. Expression of IRBP increases from E13.5 to E20.5, when expression plateaus at E20.5 and adult levels are seen [35]. To label cells that are still proliferating at P2, mice were injected with Bromodeoxyuridine (BrdU). IRBP and BrdU co-localization demonstrated that 40% of the proliferating cells at this age synthesize IRBP protein [35]. Using in situ hybridization, IRBP gene expression is detected in the retina very early in development at E10.5, with stronger expression seen at E13.5 [37]. At later stages in development, when the retinal cells have differentiated, IRBP expression is localized only to the photoreceptor cells. Unexpectedly, IRBP expression was seen in the eye lens at E10.5, with the signal disappearing after E15.5. It is not known at this time whether the disappearance of the signal is due to a masking of the signal by high levels of crystallin gene expression or if IRBP expression is shut down. IRBP expression is seen in the prenatal pineal gland and possibly in the prenatal ciliary body as well [37]. The two different techniques reveal a delay in developmental timing of IRBP expression: Western analyses show IRBP expression at E17 but not E15, whereas rtPCR and in situ hybridization show IRBP expression as early as E10.5. This difference could be due to the increased sensitivity of the rtPCR and in situ hybridization techniques or a delay in the synthesis and accumulation of IRBP protein. As was seen in mice, IRBP expression in humans is present early in development. Using complementary DNA (cDNA) from whole retina as a template in rtPCR experiments, a faint rtPCR product for IRBP is seen at fetal week 10.5 (Fwk 10.5) but not at Fwk 9.5, indicating that the onset of IRBP expression is between Fwk 9.5 and 10.5 [36]. In similar experiments done by another group, rtPCR analysis did not detect IRBP expression at Fwk 10, but cone rod homeobox gene (Crx) expression was detected at this time. Later in human eye development, at Fwk 12, IRBP, Crx, and Nrl expression was detected by rtPCR [38]. (More information about Crx expression is found in a later section). In humans, retinal differentiation first occurs in the fovea and moves in a central-toperipheral wave through the retina [38]. To determine the earliest time of IRBP expression and the specific cellular localization of the IRBP protein, immunocytochemistry (ICC)

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was performed on human eyes specifically looking at the fovea. ICC detects IRBP protein in some foveal cone precursor cells by Fwk 9. NRL-positive nuclei were first found in the fovea at Fwk 11 [38]. At Fwk 12, IRBP protein is still detected only in the fovea and now is seen in the apical region of the fovea cone precursors as a continuous band of immunoreactivity, indicating that many of the cells are expressing IRBP [39]. rtPCR of cDNA from whole eyes underestimates the time of onset of expression of IRBP but accurately recapitulates the order of expression. IRBP is expressed early in retinal development in cone precursors and is one of the earliest photoreceptor-specific proteins to be expressed. VARIABILITY IN IRBP EXPRESSION Conditions under which animals are raised and maintained also affect IRBP expression. Kutty et al. [40] demonstrated that ambient lighting conditions play a role in the regulation of IRBP but not S-antigen gene expression. Dark-reared mice showed a marked decrease in IRBP messenger RNA (mRNA) levels when compared to control animals. The dark-reared animals showed normal morphological development and normal distribution of IRBP protein. Light deprivation had no effect on IRBP protein levels, yet IRBP mRNA levels were decreased in these animals. The mechanism of light regulation on IRBP expression is unknown and could be at the level of gene transcription, RNA stability, or protein degradation. Dietary intake of retinoids can also affect IRBP expression. Katz et al. [41] fed animals a diet containing vitamin A in the form of retinyl palmitate (+A) or retinoic acid (−A). It is known that the −A diet does not support visual function but can satisfy the vitamin A metabolic requirements of most other tissues. Retinal degenerative changes occurred in the animals fed the −A diet, and decreased levels of IRBP were also found in retinoiddeprived animals (animals on the −A diet). However, the localization of IRBP immunoreactivity was not affected in these animals. This work raises the possibility that the IRBP content of the retina is regulated by retinoids that are functioning in the visual cycle. MOLECULAR BIOLOGY OF IRBP The DNA sequence for IRBP reveals an internal quadruplication in the IRBP protein structure [42, 43]. Northern blot analyses using an IRBP cDNA clone as a probe have shown that the mRNA size varies depending on the species, approximately 8 kb (bovine [44]) or 4.4 kb (human [45]). IRBP mRNA has been detected in a number of animal orders, including primates, ungulates, rodents, lagomorphs [46], amphibians [47], chickens [48], and fish [49–51]. Northern blot analyses have also been performed to demonstrate IRBP tissue-specific expression. IRBP mRNA has been found in the retina and pineal gland but not in the lens or the liver [44, 52]. Surprisingly, there have been no reports on multiple-tissue Northern blots that were probed with IRBP to search for other tissues that might express this protein. In situ hybridization of selected organs has localized IRBP mRNA to the photoreceptor cells and pinealocytes [53]. IRBP is expressed in both rods and cones [54]. Recent searches of the Gene Expression Omnibus (GEO) database (http://www.ncbi.nlm.nih.gov/ geo/) and related microarray resources suggest

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that the bulk of expression is in the pineal and retina, with low-to-nonexistent levels elsewhere. The exception is some expression in the thymus of some mouse strains [55]. There are case reports of IRBP expression in cultured cell lines (e.g., 17440 Thy+ cells), the significance of which is unknown, and numerous accounts of IRBP expression in cultured retinoblastoma cell lines, as would be expected [56]. IRBP GENOMIC CLONING We [43] were the first to describe the complete sequence of the IRBP gene. The bovine IRBP gene is compact considering the large size of its mRNA; it contains only four exons, two of which are large. The deduced amino acid sequence indicates that the protein contains a leader sequence, which would be expected in a secreted protein. We hypothesized that the unusual gene structure and fourfold repeat is the result of gene duplication and intron loss. Our group mapped the CAP site, the start of transcription, to denote where the promoter region of the gene and the gene itself begin. Other groups deduced the sequence for the human IRBP gene [45, 57]. Subsequently, we determined the sequence of the mouse gene [58]. More recently, the genomes of several vertebrates have been determined; as a result the IRBP gene structure is known in those species. These structures are discussed in more detail in the section Evolution of IRBP. Comparison of the nucleotide sequences of the 5′ flanking region from mouse, bovine, and human reveals sequence similarities between all three species in two places: upstream from the CAP site for about 300 bp, the proximal promoter region, and another 260-bp stretch that is upstream a further 1.2 kb, the distal promoter region [59]. Although these IRBP orthologs do not contain a canonical TATA or CAAT box, the nucleotide sequence around the CAP site matches the consensus sequence for a transcriptional initiator, Py Py A+1 N T/A Py Py [60]. EVOLUTION OF IRBP Because of IRBP’s location in the space between the photoreceptor outer segments and the RPE, IRBP might be necessary to maintain retinoid isomerization and chemical forms while retinoids cross back and forth [1, 61]. This hypothesis is consistent with the absence of IRBP in the invertebrates and the entirely different way that invertebrates reisomerize all-trans retinal back to 11-cis retinal while still bound to opsin in the photoreceptor cell. The mammalian IRBP gene includes a remarkable quadruplication of a 300-amino acid long repeat or module [58], while the zebrafish IRBP gene appeared to contain only two repeated modules [49]. These two different but related gene structures were puzzling. We recently expanded our search for variations in IRBP that might rectify those observed structures with prior prediction of a common ancestral gene, which was hypothesized to have appeared coincident with the vertebrate eye. We examined the IRBP gene ortholog from seven mammals: Xenopus tropicalis, the chicken, and four teleost fish. We also looked for the IRBP gene in urochordates, but we could not find an ortholog. The tetrapod IRBP gene retains a single common gene structure, with a large first exon encoding three full repeats and parts of the fourth and final repeat spread among the first through fourth exons (Fig. 1).

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Fig. 1. The tetrapod interphotoreceptor retinoid-binding protein (IRBP) gene. Orthologs of the IRBP gene are illustrated. A Mammals; B a bird and an amphibian. The same pattern of exons and introns are found in all these orthologs. Exon 1 encodes about 1,000 amino acids of the 1,200 that are normally found in IRBP and encodes three full repeats. The remainder of the protein, which includes only repeat 4, is split among exons 2–4. The positions of the introns are virtually identical in all the orthologs. (Reprinted from [119].)

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Fig. 2. The interphotoreceptor retinoid-binding protein (IRBP) gene locus in four teleost fish. The complete genome sequences have been obtained for four fish: zebrafish (A), tetraodon (B), fugu (C), and medaka (D). Each of these fish contains a single locus that contains all the IRBP gene paralogs in each whole genome. In one case, the locus contains only one gene (medaka), in a second species, there is one full copy of an IRBP gene and a small fragment of an additional gene (tetraodon), and in two species, there are two IRBP genes in the single locus (zebrafish and fugu). In the latter case, the two genes are oriented head to tail. Also, the two paralogous genes of the single locus are quite different in gene structure. The first gene contains only a single exon, which resembles exon 1 of the tetrapod gene (as though the remaining three exons were deleted). Gene 2 has four exons like the tetrapod gene, except the two internal repeats have been deleted from exon 1. (Reprinted from [119].)

The teleost fish IRBP gene locus exhibited marked differences from the tetrapods. The fish locus possesses a single IRBP gene in the Japanese rice fish (medaka; Orizias laptipes). Yet in the other three species (zebrafish, fugu, and tetraodon), there was clear evidence of an additional gene, or a remnant, upstream of the ortholog of the rice fish IRBP gene (Fig. 2). We were surprised to detect this upstream paralog, but its gene structure is illuminating. The upstream gene, in its full form, includes the first three repeats of the tetrapod IRBP gene, while the teleost fish second gene includes only the first and fourth repeats. Each teleost gene structure contains the pattern of exons that are predicted based on the content of the specified repeats from the tetrapod gene. This provides compelling evidence that the apparently unusual gene loci in the teleost fish all arose from a common ancestral gene that resembles the tetrapod gene in structure. This is illustrated in Fig. 3 along with a model of the evolution of the IRBP gene. We suggest that the IRBP gene arose about the same time as the vertebrates. We expect that IRBP arose by exaptation of another superfamily member of the IRBP protein family, which includes numerous members, all of which contain a similar three-dimensional fold

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Fig. 3. A model for the evolution of the interphotoreceptor retinoid-binding protein (IRBP) gene. The IRBP gene likely arose when the vertebrates diverged from the urochordates. This is the time period when nature “invented” the neural crest and the skull. The intent in this model is to highlight that the proposed early internal quadruplication of the IRBP gene occurred shortly after the vertebrate diverged from the urochordates (which lack the IRBP gene). This model is based on the work of Borst et al. [43] and Rajendran et al. [49]. The major revision is the addition of a two-gene IRBP locus in the teleosts. We propose that the teleost IRBP genes were formed as a result of the combination of an early whole-genome duplication and, following a short period of drift, compaction by unequal recombination to produce a two-gene locus in head-to-tail orientation [51]. A second possibility is a single-gene tandem duplication with subsequent drift of the paralogs [51]. (Reprinted from [119].)

with a substrate-binding site for hydrophobic substrates or ligands. We predict that this gene rapidly quadruplicated itself internally. Because the urochordates and the invertebrates all lack the IRBP gene, this pins the earliest appearance of IRBP to the vertebrates. This could be tested by further examination of sighted invertebrates, especially those having ciliate-type photoreceptors or intermediates having both ciliate- and rhabdomeric-type photoreceptors. Scallops bearing numerous eyes might be an appropriate test case. This model predicts that shark rays and cartilaginous fish should possess a tetrapod-like fourrepeat, four-exon gene. The elephant shark whole-genome project is being completed and might reveal informative data in the near future. The genome and IRBP sequences from nonteleost bony fish such as gars and bowfin might be informative: We predict that they will possess a single IRBP gene much like the tetrapod structure. In summary, the origins and evolution of the IRBP gene appear to coincide with the emergence of the vertebrates and the vertebrate style of eye. This coincidence may reflect fundamental changes needed with the inversion of the photoreceptor layer

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between vertebrates and invertebrates and the development of the choroid and RPE as a source of nutrients for the outer layers of the retina. This inversion of layers may in part account for the Darwinian success of the vertebrates in terms of evolutionary advantage offered by a sensory system having heightened visual acuity and improved pattern and motion detection systems. A final comment on the extraordinary teleost fish is needed: The teleost fish underwent a whole-genome duplication at about the time they diverged from other bony fish [62]. The resulting excess genetic material was freed from selective pressures, allowing three potential scenarios for each and every duplicate copy: deletion, neofunctionalization, or subfunction partitioning [63–65]. The term neofunctionalization simply represents the divergence of one of a pair of duplicate genes, one retaining the parent gene’s function while the other duplicate takes on a new and different function. Subfunction partitioning occurs when one of the daughter duplicates takes on one part of the parent’s gene functions, while the other daughter retains only complementary parts. Together the two genes complement each other and retain all the capabilities of the parent gene. Neofunctionalization occurred in the zebrafish IRBP locus, where one of the genes is expressed in the photoreceptor cells and the pineal gland, as expected, while the other gene is newly expressed in the inner layers of the retina [51]. A guiding principle should be recognized: The teleost fish genomes represent a huge opportunity to investigate how multidomain proteins work. There are over 20,000 teleost fish, each with about a one third chance of bearing a neofunctionalized or subfunction-partitioned gene of interest. It is now possible to find unusual gene structures by database searching of the several teleost fish whole genomes that are publicly available. These new gene structures, having different functions or subfunctions, help to assign physiological roles to any human gene that previously was refractory to analysis. IMPORTANCE OF THE STUDY OF THE CONTROL OF GENE EXPRESSION All cells with nuclei contain the same genetic information, yet cells are different from each other due to the expression of the different subsets of genes within each cell. The pattern of RNA abundance is characteristic of a particular cell type. The retina contains five major classes of neurons with specialized cell types within each class. The study of the control of gene expression begins the elucidation of what makes cells different from one and other. There are many different points at which gene expression can be controlled. The regulation of gene expression can occur at the level of transcription or translation. External signals can change the repertoire of genes expressed by a cell, as was seen by the dark rearing of animals and IRBP expression levels [40]. It was noticed in the late 1970s and early 1980s that the 5′ flanking region of mammalian genes contain conserved sequence motifs that are important in the control of transcription [66]. These DNA sequence features were initially used to build models of eukaryotic transcription based loosely on the classical works of molecular biology on bacterialpromoters, operators, and repressors, which usually are located immediately adjacent to the gene that they control [67, 68]. Elegant models of the process of eukaryotic RNA polymerase II complex formation and the initiation of RNA synthesis have since been built.

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Within the niche of vision research, there is a notion that more or less the exact same transcriptional processes occur in the human eye, but with specialized sets of transcription factors and transcription factor binding sites (TFBSs) that ensure specificity and selectivity of transcription during the development of the eye and, following that, during the required maintenance of terminally differentiated cells of the eye. Transcription factors are proteins that bind to sequence-specific regions of DNA and decipher (switch on) the genetic information that encodes the regulation of gene transcription (for review, see [66]). These proteins contain a DNA-binding module that is linked to one or more activation or repression modules. There is a variety of DNA-binding modules, including those containing helix-loop-helix (HLH), homeodomain, zinc finger, and leucine zipper motifs. Some transcription factors, such as those containing an HLH or homeodomain motif, can interact to form homo- or heterodimers. Transcription factors are members of multigene families that have expanded during evolution [69]. As each gene that is expressed in the eye was sequenced, inevitably it was compared with known conserved TFBS motifs and patterns in the hunt for the elements that might control its promoter. Computer-mediated studies of this type can make numerous cross comparisons, among them cross-species comparisons of orthologs and cross comparisons of genes that share expression patterns. From these computer-mediated studies of DNA sequences, many candidate cis elements were predicted, and numerous studies sought to test whether the predicted motifs had any effect on the promoter activity of the gene in question. Great debates have arisen over the precise nucleotide sequence of a binding site or exactly which of a series of closely related trans factors is just the right one that stimulates cell-typespecific gene expression of someone’s favorite eye protein. The net result so far has been an incomplete story, but within the story certain promoters and transcription factors are well studied. A few relationships of TFBSs and the paired protein transcription factors show some promise for adequately describing the complex behavior, interrelatedness, and coregulation that generate proper levels of the myriad transcripts needed for visual function. Initial work on the IRBP and other photoreceptor-specific promoters started in the middle-to-late 1980s and consisted of, for the most part, two joint strategies. Once sequence comparisons had been made that provided testable hypothesized cis elements, these joint experiments included first the manipulation of the 5′ flanking regions of the IRBP and other photoreceptor-specific protein genes (which included nested deletion series, point mutations, replacements, and combinations with other known cis elements) and the construction of reporter constructs. These constructs were introduced into retina cells or animals by transient transfection or creation of transgenic animals, usually mice or frogs (cf. [70]. Playing important parts in the transient transfection strategy was the implementation of permanently established cell lines such as Y79 and WERI from human retinoblastoma tumors [59, 71] and primary cultures of chick retina cells [72]. Many other variants on this approach have been used, including knockdown strategies and cotransfection of trans factor constructs for expression in cells normally devoid of any photoreceptor specific proteins. These promoter activity measurement studies have been best in the determination of the factors and sequences that stimulate or activate the promoter. They have been somewhat less successful in the measurement of circumstances that suppress or repress promoter activity. The second component of the joint strategy was to study the physical interaction of the DNA (the putative TFBS) with a protein (the putative trans factor) by electrophoretic

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mobility shift assays (EMSAs) [73], DNA footprinting assays, or numerous variants (with the chromatin immunoprecipitation [ChIP] assay a very important one). Other approaches include the yeast one-hybrid method to find proteins that interact with a putative DNA TFBS [74]. These measurements of binding interactions have been best once a TFBS has been shown to control the given promoter. The detection of legitimate, biologically significant interactions has been less successful when only a match to a consensus sequence motif has been detected. This may be explained by the relatively high backgrounds in EMSAs and too low a stringency employed in detecting matches to the consensus. Strategies to overcome these weaknesses include the use of sequence alignments among numerous species with known common visual capabilities, a strategy called phylogenetic footprinting [75]. This strategy will become much more powerful once the number of whole genomes that are completely sequenced becomes greater. Gene expression is controlled by both genetic and epigenetic elements. The genetic elements include the DNA sequences (cis-acting controlling elements) and the protein complexes that bind to the specific DNA sequences. The proteins are called trans factors and are also known as transcription factors. Some transcription factors bind DNA, and others form protein–protein interactions to act in combination and regulate expression of an individual gene. Each transcription factor may contribute to the control of many genes. There are some genetic elements that are found in every cell for basal expression and the initiation of gene transcription. Other gene regulatory proteins are not ubiquitous and are found in specific cells to either activate or repress (gene silencer) gene expression. Cell- and tissue-specific transcription factors work to control and maintain terminal differentiation. Recent work has shown that noncoding RNAs (microRNAs) also have the ability to silence activated genes. The epigenetic elements known to control gene expression include DNA methylation and chromatin structure. Histone deacetylases condense chromatin to make these regions inactive. Methylated DNA is found in transcriptionally silent regions of the genome. The IRBP gene methylation states were studied by Liou et al. [76] and Boatright et al. [77]. The study of the control of gene expression will lead to a better understanding of development, cancer, tissue-specific expression, and transcription factors. As an example, mutations in the Crx transcription factor are found in patients with retinal degeneration named cone-rod dystrophy (CORD2) [78, 79]. IDENTIFICATION OF DNA CIS-ACTING CONTROLLING ELEMENTS: IN VITRO AND IN VIVO EXPERIMENTS Determining the sequence of the IRBP gene provided the information necessary to study the mechanisms that control the expression of the IRBP gene. The regulation of IRBP gene expression has been studied using IRBP promoter/reporter gene fusion constructs. These constructs have been tested in transient transfection assays and in transgenic mice. Transient transfection assays show that −204 bp of the IRBP promoter are sufficient for gene expression in Y79 retinoblastoma cells [80]. Transgenic mice have also been used to study the regulation of IRBP expression by its promoter.

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A 1.3-kb fragment of the human IRBP promoter was shown to direct transgene expression to the retina and pineal gland in transgenic mice [81]. This same transgene expression profile was also seen using a 1.8-kb fragment of the mouse IRBP promoter [82]. In other studies, the 1.3-kb human IRBP fragment was shown to direct transgene expression to the photoreceptor cells of transgenic mice, but expression was heterogeneous throughout the retina [83]. The region required for tissue-specific expression was reduced to −212-bp of the human IRBP [84] and then to −123 bp [85, 86]. The critical region in the mouse IRBP promoter is similar in size [70]. A tissue-restricting factor is present between −156 and −70, and this region was refined further by Boatright et al. [87] to between −156 and −140 (Fig. 4 ), as illustrated by the differential expression of these promoters but not of the −70 promoter in Y79 and Neuro2a cells. This site appears to be the same one as described by Otteson et al. [88, 89] as the site to which Krüppel-like factor 15 (KLF15) binds in the human IRBP and opsin promoters. Two sequences in the IRBP promoter were identified within the −123-bp fragment of the IRBP 5′-flanking region that are important for the regulation of IRBP gene expression. They consist of the GATTAA sequence and its inverted repeat, AATTAG, just upstream (Fig. 5; [71, 85]. The region containing these two sites is protected from digestion in in vitro deoxyribonuclease I (DNase I) protection experiments and has been identified by several groups (see Fig. 5; [74, 85, 86, 90]. These different groups used different protein pools in the DNase I footprinting assays yet achieved very similar results. Bobola et al. [85] used a 147-bp fragment of the human IRBP promoter and nuclear extracts from retinoblastoma cells to identify a 14-bp footprint in this region of the IRBP promoter (Fig. 5]. Other groups [74, 86, 90] identified larger footprints of the same region (Fig. 6). Fong and Fong [90], also using a fragment of human IRBP promoter sequence and nuclear extracts from retinoblastoma cells, identified the larger footprinted section from −38 to −73 bp of the IRBP promoter, and this region binds to the protein OTx2 in a yeast one-hybrid experiment. Fei et al. [86] used a fragment of the human IRBP promoter sequence and a nuclear extract isolated from bovine retinas to identify a retinal-specific footprint between −42 and −76 bp of the IRBP promoter. Chen et al. [74], using a fragment of the bovine IRBP promoter and a recombinant fusion protein containing the Crx homeodomain, footprinted the region −29 to −81 bp. It is likely that numerous proteins bind to the DNA located in the footprinted region between −45 and −75 bp of the IRBP promoter. Possible candidates for proteins that bind to this region include Rx, Otx2, Crx, and Mok2. The AATTAG element had been previously identified in the mouse arrestin gene and named PCE I (photoreceptor conserved element I) because it binds retina-specific nuclear factors [91]. The PCE I core consensus sequence was determined to be CAATTAG. Closely related sequences are found in promoter regions of other vertebrate photoreceptor-specific genes, including the IRBP and opsin 5′-flanking regions. In the IRBP promoter, the PCE I sequence is conserved between the mouse, bovine, and human genomic sequences. The two regions in the mouse IRBP gene containing the PCE I are antisense strand between −1415 and −1397 and on sense strand between −74 and −55 upstream of the transcription start site [91].

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Cell Type Fig. 4. Suppression of interphotoreceptor retinoid-binding protein (IRBP) transcription in nonphotoreceptor neuronal cells. A small specific conserved sequence between positions −156 and −140 in the mouse IRBP 5′ flanking region is utilized to repress IRBP transcription in neuronal cells that are not photoreceptors. The left four columns illustrate high-level expression of a reporter gene, chloramphenicol acetyl transferase (CAT), in photoreceptor-like WERI-Rb1 cells, which originated from a retinoblastoma tumor and exhibit several photoreceptor-like properties. The right four columns illustrate the corresponding expression levels in Neuro2a cells, which originated from a neuroblastoma tumor and exhibit numerous neuronal-like characteristics but none that are specific to photoreceptor or light-sensitive properties. In the absence of a promoter, the base expression vector, (labeled pSVOATCAT), shows low background levels of CAT activity in each of the two cell lines. An IRBP promoter fragment from −70 to +101 (labeled p70) exhibits high CAT expression in both cell lines, demonstrating that this fragment is sufficient to promote IRBP transcription in these two neuronal cell lines, and we further demonstrated in a transgenic frog [70] that this holds true for all neuronal cell lineages. A construct containing −140 to +101 (labeled p140) exhibited the same expression pattern in the WERI and Neuro2a; however, a construct containing −156 to +101 (labeled p156) was active in WERI cells but inactive in Neuro2a cells. This pair of constructs highlights the region between −140 and −156 as a critical sequence in the IRBP 5′ flanking region that is apparently essential to prevent expression in an incorrect type of neuronal cell. We proposed that an active suppression event occurs, such as the binding of a trans factor to this sequence, to repress an otherwise active promoter. Further, we proposed that this factor would be absent or lack repressive activity in photoreceptor cells. Subsequent work by Otteson et al. [88, 89] identified KLF15 as a factor with exactly these properties when tested with the human IRBP promoter and human rhodopsin promoter. The sequence numberings of the mouse and human IRBP promoters are slightly different because of small insertions/deletions across the two orthologous sequences.

TRANSCRIPTION FACTORS AND THEIR ROLE IN THE CONTROL OF IRBP EXPRESSION Rx/rax Transcription Factor The transcription factor Rx binds to the PCE I element in vitro. Transfection of HEK293 cells with the mIRBP-1783/CAT (chloramphenicol acetyl transferase) construct, and hRX expression plasmid shows that Rx transactivates CAT expression up to

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Fig. 5. Footprint overlap comparison in the −70 to −45 region of the interphotoreceptor retinoid-binding protein (IRBP) 5′ flanking sequences. This sequence is identical in the human and mouse IRBP genes. Boxes represent the DNA sequence of the IRBP promoter protected in deoxyribonuclease (DNase) I footprinting experiments. Medium gray: This sequence is a portion of the entire footprint (bovine [74], human [86, 90]. Light gray [85]. The short, solid lines directly above or below the DNA sequence denote core consensus sites for nuclear factor binding. Dashed lines denote the sequence is on the reverse strand (1: [91]; 2: [74]; 3: [105]). The GATTAA invert repeat is in bold [71, 85].

14.8-fold in a concentration-dependent manner [92]. Rx-specific antibodies were used on immunoblot to show that Rx is present in adult mouse retina and iris but not liver, lens, or brain. Immunohistochemistry of adult rat retina shows Rx staining in ONL (nuclei of photoreceptors) (92). Rx/rax is a member of the paired-like homodomain family of transcription factors (93, 94). In the mouse, strong Rx/rax expression is found in the anterior neural plate of E8.5 embryos. By E10.5, Rx/rax expression is restricted to the developing eye and forebrain regions. At E15.5, Rx/rax is expressed uniformly in the neuroretina of the eye. In later stages of retinal development, there is a progressive reduction in Rx/rax expression in the retina and by P6.5 Rx/rax expression is found only in photoreceptor and inner nuclear layers. As cell proliferation ability decreases so does Rx expression. There is a correlation of expression of Rx/rax to the temporal and spatial patterns of mitotic activity in the retina (93, 94). By P13.5, in the mouse, Rx expression is undetectable in in situ hybridation experiments (94). However, northern blot analysis shows that Rax is expressed in the adult mouse retina (93). The differences seen in Rx/rax expression levels in the adult mouse may be due to the different probes that were used to detect Rx/rax expression. A 1.2-kb fragment that codes for almost the complete rax cDNA sequence was used in the northern blot experiments (93) while homeobox sequences were used in a 5’-RACE reaction to generate probes

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Fig. 6. Footprint overlap comparison in the −351 to −1 region of the interphotoreceptor retinoid-binding protein (IRBP) 5′ flanking sequences. The mouse IRBP 5′ proximal flanking sequence is shown and marked with protected regions that are color coded according to each indicated study [59, 76, 86, 85, 74, 89, 90]. There is a degree of concordance in that about 200 nucleotides are not protected by any form of retina-specific protein or nuclear extract, and most of the sequences that are protected are confirmed by two independent experiments. This is despite the (1) differences in sequences across three species, (2) differences among the several different sources of proteins (some very crude nuclear extracts and others highly purified individual proteins), and (3) differences in techniques and stringencies in deoxyribonuclease (DNase) digestion conditions.

used in the in situ experiments (94). Rx/rax is required for eye formation from the early stages of eye development. The initial specification of retinal cells and their proliferation is likely regulated by the Rx/rax transcription factor. NrL Transcription Factor Neural retina leucine zipper (Nrl) is a transcription factor that is preferentially expressed in rod photoreceptor cells. It contains a basic motif–leucine zipper and has a synergistic effect with Crx on rhodopsin transcription regulation. In the mouse retina, Nrl transcripts are detected at E12 [93]. Ablation of the Nrl gene in gene-targeted mice results in a complete loss of rod photoreceptor function but enhanced cone-mediated activity that is attributed to S cones. At 5 weeks of age, Nrl null mice retinas contain

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a similar number of nuclei in the ONL when compared to wild-type retinas; however, the rods have shorter outer segments with an abnormal disk morphology. Northern blot analysis shows that IRBP expression levels are unchanged between wild-type (+/+; 1.0), hemizygous (+/−; 1.7), and Nrl null (−/−; 1.5) mice at P10 [94]. This result is unexpected. In vitro transactivation assays measure the ability of a transcription factor to regulate the transcription of a reporter gene that is downstream of a promoter. HEK293 cells were transfected with an IRBP promoter-reporter gene and plasmids that express transcription factors. Although no statistically significant differences were observed in transactivation activity when cells were transfected with or without an Nrl expression plasmid, a synergistic affect on transactivation activity was seen when both Nrl and Crx expression plasmids were used in these in vitro experiments [88]. Peng and Chen [95], however, showed that Nrl does transactivate the IRBP promoter in HEK293T cells. Both groups used the same IRBP-reporter gene plasmid, bRbp3-300-luc. The different results found by these two groups may be due to different levels of recombinant protein expression that is caused by different transfection efficiencies or the use of different expression plasmid vectors or different cell lines. Peng and Chen used the HEK293T cell line, while Otteson et al. [88] used HEK293 cells. HEK293T cells contain a stably integrated copy of the SV40 large T antigen [96] that could produce higher copy numbers of the expression plasmid in the transfected cells. A higher concentration of Nrl protein may be required to elicit detectable transactivation activity of the IRBP promoter. Peng and Chen found that there is synergistic activity of Nrl with either Crx or Otx2 on the regulation by the IRBP promoter. Also, Nrl binds to a fragment of the IRBP promoter in ChIP assays, indicating that Nrl interacts with the IRBP promoter in vivo [95]. This interaction is not dependent on the Crx binding because ChIP assays show that Nrl binds to the IRBP promoter in retinas from Crx null mice [95]. Crx Transcription Factor The cone rod homeobox gene (Crx) encodes a paired-like homeodomain protein, is a member of the otd/Otx gene family, and was first identified in the rhodopsin promoter [73, 74]. Northern blot analysis for Crx shows a single, abundant message in the adult mouse retina, and this expression was not seen in any other organ that was studied [73]. Crx expression is first detected in the mouse at E10.5 by rtPCR analysis [36] and at E12.5 by in situ hybridization experiments [73]. In human retinas, the onset of Crx expression is between Fwk 9.5 and 10.5 as shown by rtPCR analysis, and expression is maintained in the adult [36, 38]. The use of other techniques shows that Crx mRNA is first detected by in situ hybridization experiment at Fwk 13, while ICC experiments first detect Crx protein 2 weeks later at Fwk 15 [36]. The difference in timing is likely due to the difference in the threshold of sensitivity between the different techniques. In vitro, Crx transactivates IRBP promoter-reporter gene constructs in a dose-dependent manner [74, 92]. Transactivation activity of the IRBP promoter is enhanced when both Crx and Nrl expression plasmids are used [95]. When the Crx site in the mouse IRBP promoter was mutated, there was a suppression of activity, but it was not abolished. The remaining transcription activation activity detected is likely due to the PCE I/Ret1 element [71]. ChIP assays show that Crx binds to the IRBP promoter in vivo [95].

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In both human and mouse retina development, the onset of expression for both Crx and IRBP mRNA is very similar. Bibb et al. [36] did not find Crx protein until Fwk 15—several weeks after the mRNA expression is detected. The developmental expression pattern and the in vitro work both suggest that Crx is a transcriptional activator of IRBP expression (positive transcription factor for IRBP). A Crx knockout mouse was created by a targeted deletion of the homeodomain region of the Crx gene [97]. At P10, before the outer segments begin to form, there is no phenotypic difference seen in retinas of the different Crx null mouse genotypes. At P14, outer segments are not seen in Crx null mouse retinas, and in Crx hemizygotes the outer segments are shorter than those of wild-type mice. The expression levels of photoreceptor-specific genes were determined in Crx null mice mutant retinas by densitometry of northern blots from P10 wild-type, hemizygous, and Crx null mice. The expression levels for IRBP are the same in all three genotypes of mice, unlike other photoreceptor-specific genes such as rhodopsin and cone arrestin, which have expression levels that are tenfold lower when the hemizygous and null phenotypes are compared [97]. Transcription factors interact with specific sequences of DNA and with other proteins to regulate gene transcription/expression and cellular differentiation. The Crx null mice were used in a study designed to identify the Crx transcriptional networks [98]. Crx gene targets were identified by microarray analysis of gene expression patterns from normal (wt) and Crx null developing retinas. Photoreceptor-specific and enriched genes were divided into two categories based on the downregulation of their expression in the absence of Crx expression: genes that are downregulated more than 2.5-fold or genes downregulated less than 2.5-fold. IRBP is in the second category of genes, and this agrees with earlier work that the knockout of Crx does not have a large effect on the expression of IRBP [97]. In addition, comparison of the upstream sequences from genes that were downregulated in the absence of Crx led to the identification of a 11-base sequence motif that contains one strong Crx-binding element (CBE) upstream of a second, weaker CBE-like sequence [98]. The IRBP promoter contains two GATTA sequence motifs that are slightly similar to the CBE motif (Fig. 5). It contains the second weaker CBE-like sequence but not the first, stronger element, suggesting that Crx is not a main contributor to the activation of the IRBP gene. These two GATTA motifs are required for IRBP expression because mutations in both GATTA motifs, found in the −58 to −45 bp region and the second one just upstream (inverted), abolish expression of the reporter gene in transgenic mice photoreceptors [85]. The regulation by Crx may not contribute significantly to the overall transcriptional regulation of the IRBP gene, or the expression of IRBP may be regulated or coregulated by Otx2 because the consensus sequences for Crx and Otx2 are similar. Crx and Otx2 have an 86% homology in the homeodomain [99]. In Crx null mice, Otx2 activity may compensate for Crx activity in the regulation of IRBP gene expression, thus resulting in a minimal change in IRBP expression levels in these mice. Crx antisense knockdown suppresses p70 activity, suggesting that whereas Crx is not required for IRBP expression, it may in fact be used endogenously [100]. OTX2 Transcription Factor Otx2 is another paired-type homeodomain protein, a member of the Otx homeobox gene family, and is expressed in the retina [101]. Otx2 is an essential developmental gene. It has

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multiple roles in gastrulation and brain and retinal development. A targeted disruption of the Otx2 gene leads to embryonic lethality at midgestation, thus preventing the analysis of the influence of Otx2 expression retina development [102]. Conditional knockout mice show that Otx2 is required for neuronal specification of the retina and for pineal gland development. Absence of Otx2 results in the conversion of differentiating photoreceptor cells into amacrine-like neurons and to deficiency of pinealocytes in the pineal gland [103]. In adult mice, Otx2 immunoreactivity is found in several retinal cell types, including the nuclei of photoreceptors [99] or the cytoplasm of rod photoreceptors [101]. The subcellular location of the Otx2 protein appears to be antibody specific. In situ hybridization experiments studying Otx2 expression in the developing mouse retina show that at E11.5 strong staining is seen in the RPE, while staining is weak in the neural retina [103]. At E12.5, expression of Otx2 in the neural retina has increased, and by E17.5 there is very strong staining for Otx2 in the outer region of the neuroblastic layer. At P6 and in the adult, faint staining is detected in the ONL, suggesting that there is weak expression of Otx2 in adult photoreceptor cells [103, 104]. This is in contrast to Crx (another member of the Otx gene family) expression. In the adult mouse retina, strong staining for Crx expression is found in the ONL [103]. Otx2 was identified as binding to the IRBP promoter in a yeast one-hybrid screen using a human retina library and an IRBP sequence that was protected in DNA protection experiments [90]. The Otx2 protein transactivates the IRBP promoter in vitro. When HeLa cells are cotransfected with a 123-bp hIRBP promoter-reporter gene construct and an Otx2 expression plasmid, IRBP transcription is activated five- to sevenfold [99]. Cotransfection of the hIRBP reporter gene construct and a Crx expression plasmid increases IRBP promoter activity four- to fivefold, which is in agreement with the transactivation experiments of Chen et al. [74] in HEK293 cells. Transfection of HeLa cells with the 123-bp hIRBP reporter gene construct, Otx2 expression plasmid, and Crx expression plasmid does not increase IRBP promoter activity, suggesting that the Otx2 and Crx proteins do not act synergistically on the human IRBP promoter [99]. Otx2mediated transactivation of the IRBP promoter was also seen when a plasmid containing the region −66 to +68 bp of the IRBP promoter and a reporter gene was cotransfected with an Otx2 expression plasmid into HeLa cells [90]. This same plasmid was used to study the transactivation activity of different isoforms of Otx2 in 293T cells [104]. In these experiments, the B isoform had stronger stimulation of the IRBP promoter than the A or C isoforms; this was attributed to higher levels of translation of the B isoform of Otx2 in this in vitro model [104]. Transactivation activity of the IRBP promoter is enhanced when both Otx2 and Nrl expression plasmids are used [95]. Also, Otx2 binds to a fragment of the IRBP promoter in ChIP assays, indicating that Otx2 interacts with the IRBP promoter in vivo [95]. ChIP assays also show that Otx2 binds to the IRBP promoter in retinas from Crx null mice, so the interaction between the IRBP promoter and Otx2 is not dependent on Crx binding [95]. Otx2 is expressed before Crx in mouse retinal development. Experiments in which the Otx2 genes are inactivated under the control of the Crx promoter (in conditional knockout mice) show that Otx2 is a direct regulator of Crx gene expression [103]. The above-listed trans factors are some of those needed to achieve cell-type selectivity and specificity of expression in the retina. It is widely recognized that the rod photoreceptor [93] does express many other trans factors, as is indicated by microarray

106

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studies. Given this plethora of trans factors, it is not yet possible to predict when the story of IRBP transcription will be complete. TRANSGENIC MICE Transgenic mice were generated using the hIRBP promoter fragment −123 to +18 upstream of a reporter gene. When both the PCE I and CRX elements are mutated in transgenic mice, reporter gene activity is abolished in photoreceptor cells [99]. In vitro experiments show that mutation of either the Ret-1/PCE I or the CRX elements suppresses IRBP promoter activity [71]. The same fragment of the hIRBP promoter (−123 to +19) upstream of a reporter gene was used to make transgenic mice, and either the PCE I element or the CRX element was mutated [86]. Mutation of the CRX but not PCE I element abolishes reporter gene activity in photoreceptor cells. When the PCE I element but not the CRX element is mutated in the IRBP promoter, 9 of 17 lines of transgenic mice show photoreceptor-specific expression of the reporter gene [86]. These studies indicate that the CRX DNA element is required for photoreceptor-specific expression of IRBP. rtPCR experiments, using the different lines of transgenic mice, show that to have IRBP promoter activation, a protein interaction with the PCE I element is not required but is enhanced when the PCE I element is present [86]. REPRESSORS OF IRBP GENE EXPRESSION A potential strong silencer was identified in the region between −206 and −66 bp of the mouse IRBP promoter in transient transfection experiments using nested deletions of the 5′-flanking region controlling expression of a reporter gene [90]. KLF15 Krüppel-like factor 15 (KLF15) was identified in a yeast one-hybrid assay using a bovine retinal cDNA library and a 29-bp fragment of the bovine rhodopsin promoter as bait. Transactivation experiments were performed in vitro to study the activity of KLF15 on the IRBP promoter since the rhodopsin and IRBP DNA sequences share some gene regulatory elements [88]. HEK293 cells were transiently transfected with a KLF15 expression plasmid and an IRBP promoter-reporter gene construct containing the proximal region of the bovine IRBP gene, −300 to +132 bp. Expression of KLF15 reduced reporter gene activity of the IRBP-reporter gene construct in a dose-dependent manner [88]. CRX expression is known to transactivate the IRBP promoter in vitro [74]. Reporter gene activity was also repressed by KLF15 expression when CRX or CRX and NRL were cotransfected with the IRBP-reporter gene construct and the KLF15 expression plasmid [88]. The zinc finger domains of the KLF15 transcription factor bind to the IRBP promoter [89]. DNA footprinting experiments using a KLF15 fusion protein identified three KLF15 binding sites in the bovine IRBP promoter. The same footprints are found on both positive and negative strands and are conserved among three species. DNA binding is zinc dependent, and there is evidence that retinal proteins bind to the KLF15 site. The footprinted site, K1-c, is next to the CpG dinucleotide (−115) that is hypomethylated specifically in retinal cells

IRBP Molecular Biology

107

[35, 77]. Perhaps KLF15 is regulating the methylation/chromatin structure of the IRBP 5′-flanking region for the repression of IRBP expression in the inner retina. MOK2 MOK2 is a Krüppel/TFIIIA-related zinc finger protein. MOK2-binding sites are present in the IRBP gene, and experiments were performed to determine if IRBP is a potential MOK2 target gene [105]. In the IRBP promoter, 8 bp of the MOK2 core sequence is found (TTAAGGCT) in the reverse orientation, and this overlaps the CRX/ OTX2 site (Fig. 5). The IRBP DNA sequence defined as the MOK2 site is conserved between human, bovine, and mouse IRBP promoter sequences. A MOK2-binding site is also found in intron 2 of the human and bovine IRBP genes [105]. The IRBP promoter region binds to MOK2 recombinant protein in EMSAs, and the band is supershifted with an anti-MOK2 antibody showing that MOK2 protein binds to the IRBP promoter in vitro [105]. In transient transfection experiments of Weri-RB1 cells with an IRBP promoter-reporter construct alone or with MOK2 expression plasmids, a significant reduction in transcription activity was observed when the MOK2 protein was present, showing that MOK2 can act as a transcriptional repressor of the IRBP promoter activity in vitro. However, the sensitive technique of in situ rtPCR was required to detect a low level of Mok2 expression in a subset of nuclei in the ONL of 1-month-old mouse retinas [105]. Chx10 Chx10 gene expression is required for retinal progenitor cell proliferation and bipolar cell differentiation. Chx10 is required to block rod cell differentiation but is not essential for the proliferation of progenitor cells in the postnatal retina [106]. It contains a pairedlike homeodomain [107]. Chx10 represses transcription in vitro [108]. Chx10 knockdown inhibits the generation of bipolar cells and differentiation of rod photoreceptor cells. When overexpressed in newborn mouse retinas, Chx10 promotes the differentiation of bipolar cells without affecting cell division or survival [106]. The expression of Chx10 is found early in retinal development in the mouse at E9.5 in the outermost region of the evaginating optic vesicle, the area that will form the neuroretina later in development [107]. At E16.5, when lamination of retina is starting and differentiated ganglion cells have been formed, Chx10 expression is detected in the neuroblast layer but not in the ganglion cell layer (GCL). Expression of Chx10 at P4 is further restricted to the future inner nuclear layer and is not found in the future ONL [107]. In the adult mouse retina, Chx10 expression is found in bipolar cells and a subset of Müller cells [109]. In the ocular retardation mouse mutant (Chx10or-J/or-J), ocular development is abnormal. The phenotype is characterized by blindness, small eyes (micropthalmia), abnormal photoreceptors, and absent optic nerve. Analysis of expression of other known transcription factors in the retina was performed to study the effect the absence of Chx10 has on these proteins. Unlike wild-type mice, Crx expression was not detected during retinal development in the embryo but was expressed after birth [110]. rtPCR experiments were performed to study the expression of proteins that are putative targets of Crx, and expression of many of these proteins was delayed in the Chx10or-J/or-J retina. The expression of

108

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IRBP was not delayed in the Chx10or-J/or-J retina [110], further supporting the idea that Crx expression is not required for IRBP expression in the embryonic retina. ChIP assays were used to identify potential gene targets of Chx10 in the in vivo chromatin context [111]. Mouse retina samples were obtained from mice at different ages and treated with formaldehyde to cross-link the proteins to their DNA targets and to each other. Sonication was performed to cut the DNA into an average size of about 1 kb. After incubation with anti-Chx-10 antibodies, the complexes were immunoprecipitated, purified, and the cross-links reversed. Three regions of the IRBP promoter were specifically chosen (targeted) for amplification by PCR to determine if Chx10 binds to the IRBP promoter at these sites. These sites were chosen because they are known to contain homeodomain-binding sites [91]. ChIP shows that Chx10 binds (in vivo) to one site in the IRBP promoter, upstream at −1.4 kb. This result was unexpected because the proximal IRBP promoter contains a PCE-1 element that is also present in the arrestin promoter. The PCE I element in the arrestin promoter does bind Chx10 both in vitro and in vivo, as shown by ChIP assays. The ChIP assays show that protein binding to DNA is dependent on chromatin context. Chx10 binds IRBP promotes in vivo at P0, P6, and P14. The consensus TAATtgac that they identify is part of the DNA fragment identified by ChIPeven though this sequence is outside of the fragment that was PCR amplified [111]. These ChIP experiments are not conclusive because Chx10 could be binding to regions of the IRBP promoter that were not identified and amplified by PCR. Chx10 may be repressing IRBP gene expression in bipolar and Müller cells, but it is not acting by itself. Other factors are at work. IRBP gene expression is controlled by a combination of factors that include positive and negative regulators. The IRBP gene is not expressed in bipolar and Müller cells because repressors and not activators of IRBP expression are found in these cells. Needless to say, nature can often switch a repressor to an activator, and vice versa, with posttranslational modifications. It is worth considering potentially analogous systems that use closely related or the same proteins. In muscle, myogenesis occurs when myoblasts differentiate into myotubes. An analogous set of proteins, including homeodomain proteins and basic helix-loop-helix (bHLH) proteins can form highly coordinated and synergistic complexes to activate transcription of genes needed to build myofibers. Figures 7 and 8, which show the IRBP 5′ flanking region, illustrate that matches to consensus trans factor binding sites are located upstream of the IRBP gene in numerous species. In particular, two such sites, a COMP1 and a Chx10 match, are found about 1,200 nucleotides upstream of the transcription initiator site in a well-conserved region (hereafter called the upstream conserved region). These sites are about 65 nucleotides apart. Chx10 is well known to the vision sciences community, but COMP1, which stands for COoperates with Myogenic Proteins 1, is a poorly understood cis element [112]. This element is known to bind trans factors cooperatively and to synergistically enhance transcription of a reporter gene when transfected into myoblasts that are differentiating into myotubes. Since this trans factor binding site was first recognized in 1992, it was found [113] that the COMP1 cis-element sequence TGATTGAC can be bound in a cooperative manner by a heterodimer of PBX/Meis1-Prep1 with another heterodimer consisting of E2a bound to MyoD, myogenin, Mrf-4, or Myf-5 (PBX/Meis1-Prep1 are homeodomain proteins whereas E2a, MyoD, myogenin, Mrf-4 and Myf-5 are bHLH proteins). While it is not clear that any of these proteins are available in the adult retina or actually bind to the

B

A

RBP3

48005000

47900000

48000000

ANXA8L1

Fig. 7. (continued)

ost Conserved

Conservation

V$CHX10_01 V$BRN2_01 V$NCX_01 V$BRACH_01 V$COMP1_01 V$CHX10_01 V$ER_Q6 V$COMP1_01 V$HOX13_01 V$USF_C V$YY1_02 V$GRE_C V$GATA1_05 V$PAX5_01 V$RP58_01 V$PAX4_03 V$OCT_C

chr10:

Most Conserved

Conservation

FBS Conserved

chr10:

48000000

48050000

48020000

RefSeq Genes

48015000

48025000

48030000

12 3

4

PhastCons Conserved Elements, 17-way Vertebrate Multiz Alignment

Vertebrate Multiz Alignment & Conservation (17 Species)

HMR Conserved Transcription Factor Binding Sites

48010000

PhastCons Conserved Elements, 17-way Vertebrate Multiz Alignment

Vertebrate Multiz Alignment & Conservation (17 Species)

RefSeq Genes ZNF488 GDF2 RBP3 GDF10 HMR Conserved Transcription Factor Binding Sites

47950000

GDF2

48035000

4810000 0

IRBP Molecular Biology 109

48012380

48012390

48012400

48012420

48012430

48012440

48012450

48012460

48012470

48012480

48012490

48012500

48012510

48012520

48012530

7 1 4 A GG T C T GGGGC T A A A C T C C T G A G T T GGGGC A A GGC T T C C A GC T C C A G T A A GC C T T T A A T C C T G T C T A A T T C A A GC A C A T C A A C C C T GGG T A T CGGGG A GG A G T GGC C A GGG T GG T T T G A C C C A G A A GG T A GG T C T GGGGC T C A A T G T GGG A G T T A G A G T G A GGC T T C C A GC T C T GC T A A GC C T T T A A T C C T G T C T A A T T C A A GC A C A T G A GC T - - - - GG A C T C T G A A G A A G A GGC C A GGG T GG T T T G A C C CGG A A GG A A GG T C T GGGGC T C A A CG T GGG A G T T A G A G T G A GGC T T C C A GC T C T GG T A A GC C T T T A A T C C T G T C T A A T T C A A GCGC A T G A A C C A - GG A G A C T C A G A A G A A G A GGC C A GGG T A G T T T G A C C CGG A A GG A A GG T C T GGGGC T A A A C T CGGG A G T T GGGG T GCGGC T T GC A GC T C C A G T A A GC C T T T A A T C C CG T C T A A T T C A A GCGCG T C A GC C C C GGGG A C CGGGG A GC A GGGGC C G CGGCGG T T T G A C C C A G A A GG C A A GC C T T A T GC T A A A T A T GGG A GC A GGGGG A A GGC T T T T A T C T C C A T T A A A C C T T T A A T C T C A T T A A A T T G A A G T GC A T C A GC C - - - - - - - - - - - - A A A GGG T G A T A A T GG T GGC C A G T C C T GC T G T T T A A T C T C T GC A T A G A A T A CGGG T GC A C CG A A A A A T C T T T T A T C T GC A T T A A A C C T T T A A T C T C A T T A A A A T GG A G T GC A T C A T C C - - - - - - - - - - - - - - A A A G - - - - - - - - - - - - - - T GC A C C A A T A G A T ===================== = ==================== = ===================== = =================== = = ===================== = ==================== = ===================== = ==================== = ===================== = ==================== = ==================== = = ==================== = PhastCons Conserved Elements, 17-way Vertebrate Multiz Alignment

48012410

T GC A GC T GC C T T C CGC C C T T G T C C T T C T C A GC T GG T GG A C A G A T GC A GC T GC C T C T C T C C C T T G C C C T T C T T A GC A GGC A G A C A G A T GC A GC T GC C T T T C T C T G T T G C C C T T C T T A GC A GGC A G A C A G A T G T GGC CGC C T T C C T C C C T CG T C C T T C T C A GCGGG T GG A C A G A - - - - GC TGTGTGT CCC T C T T - - - TGT C T A TGAC T A T AGACAG A = = = = GC TGT C - - - - - - - - - - - - CC TGT T TGTGA A T CAC AC TG A ===================== = ==================== = - - - GGC T GC C - - CGG T C T T T G T = = = = = = = = = = = = = = = = = = = = =

HMR Conserved Transcription Factor Binding Sites Vertebrate Multiz Alignment & Conservation (17 Species)

AT C TGC TC AG T CAG T T AG T CA A GC A T C CGC T C == = == = == == = == = == == = == = == == = == = ==

T T T T = = = =

TT CT CT CT == == == ==

GGA A A T C GGA A GC C GGA A A T C G GG A A T C = == = == = = == = == = = == = == = = == = == =

AT AT AT AT == == == ==

T T T T = = = =

AT AT AT AT == == == ==

C C C C = = = =

T G A GG A TG A CG A TGA AGA T G G GG A == = == = == = == = == = == = == = == =

TGT TGT TGT TGT == = == = == = == =

T T T T = = = =

T T T T = = = =

A CC A A CC A A CC A A CC A = == = = == = = == = = == =

AC T GC T A G T GG T AG T AG T A G T GG T == = == = == = == = == = == = == = == =

T T T T = = = =

T T T T = = = =

A TGA A GG A A CG A A TGA = == = = == = = == = = == =

AGGCC A AGGCC A AGGCC A AGGCC A == = == = == = == = == = == = == = == =

AAGAGG TC A AGGAGG TC A AGGAGG TC A AAGAGG TC A == = == = == = == = == = == = == = == = == = == = == = == =

A T T AGC T A A AA C AA A C A T A T T T T T AGCT C A T T AGGA T A T T AGC T A A AA C AA A C A T A T T T T T AGCT C A T T AGGA T A T T A G C T A A A A C A A A C A T A T C T T T A G C T C A C T GG G A T A T T AGC T A A AA C AA A C A T A T T T T T AGCT C A T T AGGA T = = = = = = = = = = = = = = = = = = A T TC T T AGCT C A T T AGGA T == = == = == = == = == = = = == = == = == = == = == = == = == = = = = = = = = = = = = = = = = = = = = T T C T AGGT C A T T TGGA T == = == = == = == = == = = = == = == = == = == = == = == = == PhastCons Conserved Elements, 17-way Vertebrate Multiz Alignment

T T T T T = T =

2 4 TGTGT TGT T T TGT T T TGT T T TGT T T == = == TGT T T == = ==

2 A ATGCT CAAGTGACA T A A T A CCC AAG TGA CA T A A T A CCCGAG TGA CA T A A T GCCC AAG TGA CA T A A T GCCC T AG TGA CA T = == = == = == = == = == = T AT CCT AGTGT T ACA T = == = == = == = == = == =

AT AT AT AT AT == AT ==

GGT GGT GGT GGT GGT = == GCT = ==

CACA CTCA CTCA CCC A GATG = == = CACA = == =

TC TC TC TC TC == TC ==

A C T C CA C CG T A CCCCA C TG T A CC C CGC TG T ACT CCAC TGT T CCCCA C TG T = == = == = == = T GA CCA T TCC = == = == = == =

A T T T T = T =

T GCAGCC A T GCAGCC A T GCAGCC A T GCAGCC A T T CA A CC A = = == = == = T A CA A CC A = = == = == =

1 A T GGA AGT GAG A T GGA A A T GAG A T GGA A AC A AG A T - GAGGT GAG A T GGA A A T GAG == = == = == = = = A T T GA A AGC AG == = == = == = = =

Fig. 7. Human interphotoreceptor retinoid-binding protein (IRBP) gene 5′ flanking region and locus. A The locus containing the IRBP gene and adjacent bounding genes spans from roughly nucleotide positions 47,800,000 to about 48,100,000 and contains the Annexin 8 and Znf488 genes on the proximal side and the GDF2 and GFD10 genes on the distal side. GDF10, GDF2, and RBP3 (the IRBP gene) are transcribed in the same head-to-tail orientation (from right to left), while the Annexin 8 and ZNF488 genes are transcribed in the opposite orientation. B A close-up of the DNA sequence that spans from the beginning of the GFD2 gene through the IRBP gene and the positions of HMR conserved transcription factor binding sites (TFBSs) are indicated. In the two tracks below that, conserved sequences are noted.

lod=133

Gaps human mouse rat dog opossum chicken x_tropicalis tetraodon

Conservation

chr10: ---> A T C T G C T C T T T G G A A A T C A T T A T C T G A G G A T G T T T A C C A A C T G C T T T A T G A A G G C C A A A G A G G T C A A T T A G C T A A A A C A A A C A T A T T T T T A G C T C A T T A G G A T T T G T G T A A T G C T C A A G T G A C A T A T G G T C A C A T C A C T C C A C C G T A T G C A G C C A A T G G A A G T G A G RefSeq Genes RefSeq Genes HMR Conserved Transcription Factor Binding Sites V$COMP1_01 V$CHX10_01 Vertebrate Multiz Alignment & Conservation (17 Species)

2

lod=45

Gaps human mouse rat dog opossum chicken x_tropicalis tetraodon

Conservation

V$ER_Q6

RBP3 Gene

48010970 48010980 48010990 48011000 48011010 48011020 48011030 48011040 48011050 48011060 48011070 48011080 48011090 48011100 48011110 48011120 chr10: ---> T G C A G C T G C C T T C C G C C C T T G T C C T T C T C A G C T G G T G G A C A G A A G G T C T G G G G C T A A A C T C C T G A G T T G G G G C A A G G C T T C C A G C T C C A G T A A G C C T T T A A T C C T G T C T A A T T C A A G C A C A T C A A C C C T G G G T A T C G G G G A G G A G T G G C C A G G G T G G T T T G A C C C A G A A G G T RefSeq Genes

1

C

110 Borst et al.

The predictions were generated from the Multiz Alignment (which compares sequences from 17 species) and the PhastCons algorithm. Exons are well defined by these programs, but several other sequences are denoted that are strongly conserved. Four of these conserved elements are marked with arrows and numbered 1 through 4. The elements labeled 3 and 4, although strongly conserved, do not contain any currently known TFBSs. Elements 1 and 2 contain well-known TFBSs, including Chx10-binding sites. It is worth noting that GFD2 and IRBP are transcribed in the same orientation, and both genes bear TFBSs for Chx10; we speculate that these two genes might be coregulated. C Close-ups of elements 1 and 2. Element 1 is contained in the proximal IRBP promoter, and the zoom-in shows roughly 130 nucleotides of the promoter. A leftward pointing bent arrow at position 48,010,996 indicates the transcription start site. Strong sequence conservation at the transcription start site extends to either side of the start site for a few nucleotides into the gene and for about 15 nucleotides into the 5′ flanking region. Found about 50 nucleotides upstream of the transcription start site, the highly conserved element identified as “lod=45” contains the Ret-1/PCE-I- and CRX-binding sites. This sequence is almost identical in the mammals and highly conserved through the chicken, more so in the CRX binding element than in the Ret-1/PCE-I site. The latter sequence difference may reflect variation in IRBP gene expression patterns of the rod-dominant mammals compared to the cone-dominant chicken retina. Element 2 is illustrated in C2. This conserved sequence spans about 160 nucleotides and contains conserved Chx10 and COMP1 TFBSs. The work of [113] raises the possibility that cooperative binding of Meis/PBX/Prep1 may bind to these two sites with an intermediate loop in cells of the retina. This hypothesis should be testable given the availability of looping assays and antibodies that are specific to these DNA-binding proteins. It would require the isolation of photoreceptor nuclei separated cleanly from other neurons of the retina. At least two possibilities exist: the use of NRL-GFP transgenic mice [93] and cell sorting to obtain a pure population of photoreceptor cells and another of other neurons. Alternatively, it may be possible to recover chromatin from the outer nuclear layer (ONL) and other nuclear layers of the retina isolated with laser capture microscopy.

IRBP Molecular Biology 111

Lrrc18

chr14:

CpG: 19 0.1 _

0_

32775000

Mapk8

1 2

32780000

32790000

32795000

3

PhastCons Conserved Elements

Vertebrate Multiz Alignment & Conservation

ESPERR Regulatory Potential (7 species)

CpG Islands (Islands < 300 Bases are Light Green)

UCSC Known Genes Based on UniProt, RefSeq, and GenBank mRNA Rbp3 Zfp488

32785000

Vertebrate Multiz Alignment & Conservation

32800000

32805000

RefSeq Genes Gdf10 LOC432838 Ppyr1 Syt15 Glud1 Syt15 2200001I15Rik Gdf2 Antxrl Rbp3 Anxa8 3110001K24Rik Sncg Zfp488 Mmrn2 Bmpr1a

Superfamily/SCOP: Proteins Having Homologs with Known Structure/Function

Ptpn20

32810000

Ldb3 Ldb3 Ldb3 Ldb3 Ldb3 Ldb3 Ldb3 Opn4

Fig. 8. Mouse interphotoreceptor retinoid-binding protein (IRBP) locus. A The mouse IRBP gene and a region containing about 20 genes nearby from roughly position 31,800,000 to 33,400,000. The IRBP gene is bounded immediately by the same GFD2 and Znf488 genes, and there are some other genes that might be of interest, including the optineurin 4 gene and the Bmpr1a gene. Most of the identified genes possess homology with superfamilies of known proteins. An expanded view of the locus is shown in B, showing the GFD2, RBP3, and Znf488 genes. Sequence conservation illustrated in several tracks in this panel demonstrate the positions of exons in each of these three genes. Three elements indicated by the digits 1, 2, and 3 highlight sequences that are strongly conserved but found between the GDF2 and RBP3 genes. It is not clear yet whether the first two sequences represent random occurrences, whether they have a role in the termination of the GDF2 gene, or whether they function in the promoter of the RBP3 gene. The sequences indicated with “3” corresponds to element 2 in Fig. 7. This site contains matches to the consensus transcription factor binding sites (TFBSs) for COMP1 and Chx10.

Most Conserved

rat human dog opossum chicken x_tropicalis tetraodon

Conservation

Gdf2

32770000

Arhgap22

31900000 32000000 32100000 32200000 32300000 32400000 32500000 32600000 32700000 32800000 32900000 33000000 33100000 33200000 33300000 33400000

chr14:

Reg Potential 7 species

B

Conservation

Superfamil y

A

112 Borst et al.

IRBP Molecular Biology chr10:

48011000

48011100

48011200

48011300

113 48011400

48011500

48011600

48011700

48011800

48011900

48012000

48012100

48012200

48012300

48012400

48012500

48012600

GenomeTrafac-UCSC Mapping -9710-85972

Human RBP3 exons

NM_002900 1_

Alignment Curve

Concise Alignments 0.48 _

V$CP2.01:118 V$PAX5.01:610 V$PAX5.01:608 V$ELK1.02:609 V$NBRE.01:607 V$GKLF.02:606 V$OTX2.01:605 V$MEL1.01:604 V$GSH2.01:601 V$TST1.01:602 V$MSX2.01:603 V$CHR.01:600 V$ZIC2.01:599 V$ER.01:597 V$AML3.01:598 V$CHOP.01:596 V$MEL1.01:594 V$GATA1.02:595 V$CRX.01:593 V$TH1E47.01:592 V$HNF1.01:591 V$HNF1.01:590 V$XVENT2.01:373 V$CDP.01:372 V$NKX25.02:589 V$FAST1.01:371 V$CDP.01:370 V$CART1.01:369 V$GATA3.02:368 V$AREB6.04:367 V$PAX2.01:366 V$MYT1L.01:365 V$AP4.01:116 V$MYF5.01:117 V$ATF6.02:114 V$XBP1.01:115 V$DICE.01:471 V$DICE.01:470 V$AP4.01:113 V$AP4.01:112 V$COMP1.01:469 V$BARX2.01:468 V$GATA1.02:467 V$MEL1.03:466 V$FREAC2.01:465 V$VDR_RXR.01:464 V$FXRE.01:463 V$GSH2.01:461 V$S8.01:462 V$HFH3.01:459 V$CABL.01:460 V$ISL1.01:457 V$GSH2.01:458 V$DEC1.01:455 V$NKX25.01:456 V$OCT1.06:454 V$PARAXIS.01:452 V$NEUROG.01:453 V$COMP1.01:449 V$NFY.01:450 V$CAAT.01:451 V$MEL1.02:448

Conserved TF binding sites on this gene.

RefSeq Genes

RBP3

GNF Gene Expression Atlas Ratios Using Affymetrix GeneChips - Arrays Grouped By Tissue Median

GNF Ratio V$ER_Q6

HMR Conserved Transcription Factor Binding Sites

V$COMP1_01 V$CHX10_01

Fig. 9. Stringency of searching and the number of detected trans factor binding sites. Many more transcription factor binding sites (TFBSs) are detected as the search stringency is dropped. Numerous TFBSs can now be detected in addition to the high-stringency search, which identified only three conserved sites. Now, about 50 TFBSs are detected in the proximal and upstream conserved sequences of the human interphotoreceptor retinoid-binding protein (IRBP) gene. With the drop in stringency, now Otx2, Zic2, CRX, Pax2, Myf5, RxR, neurogenin, and numerous other TFBSs are also detected.

IRBP promoter (at these two imperfect matches to the Chx10 and COMP1 sites), it might be possible that related proteins form an analogous heterotetramer, perhaps even involving Chx10, that might up- or downregulate IRBP gene transcription. That said, there is clear evidence that abnormalities in Prep1 can affect Pbx and Meis1 expression and result in major phenotypic defects in the formation of the eye. Pax6 expression is concomitantly reduced [114]. Meis1-defective mice also have eye abnormalities [115]. Last, Meis1 directly interacts with Pax6 during lens formation [116]. Thus, it seems possible that these trans factors may play significant roles in the control of IRBP gene expression. Within the more proximal 5′ flanking sequences of the IRBP gene, there is only one highly conserved trans factor binding site that is detected by the HMR-conserved TFBS algorithm, and this site is an estrogen receptor alpha site (V$ER_Q6, in Transfac Database v 7.0). However, with a less-stringent set of computer parameters, about 50 trans factor binding sites can be found as illustrated in Fig. 9. Proximal to the transcription initiator site (within the first 500 nucleotides between 48,011,000 and 48,011,500), there are about 25 TFBSs, and these sites notably include Otx2, CRX, and Zic2, and all are found in the publicly available version of Transfac v7.0. The analysis of these trans factor binding sites shows few TFBSs between about positions 48,011,550 and 48,012,300, a region where homology between orthologous mouse and human sequences is limited. Just distal to 48,012,300, there is another collection of another 25 or so TFBSs in a span of about 220 nucleotides, which is the previously discussed upstream conserved region.

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This abundance of putative TFBSs highlights the remaining challenge to determine which TFBSs play a role in activating or repressing IRBP transcription. This problem is solvable, as illustrated by the important work on the very limited number of these trans factors that we discussed above. The Chx10 site between 48,012,428 and 48,012,441 in the upstream conserved region might play a role in the repression of IRBP transcription in nonphotoreceptor retinal cells. It is worth consideration of the binding properties of Chx10 in the proximal part and the upstream conserved region of the IRBP promoter [111]. It was found that in vitro in EMSAs Chx10 can bind to the Ret1/PCE I site in the proximal promoter and to the Chx10 site in the upstream conserved element. However, by in vivo CHiP assay, they found that Chx10 does not bind well to the proximal Ret1/PCE I element immediately adjacent to the CRX-binding site on the IRBP promoter. Importantly, it was found that Chx10 is strongly associated with the IRBP upstream conserved element. These findings were the same whether using mice at P0, P6, or P14. These findings may be affected by the use of whole neural retinas, as opposed to cell-type-specific chromatin (cf., from purified rod photoreceptors or highly enriched populations containing bipolar cells only). SUMMARY AND CONJECTURE Transcription factors have been identified that activate or repress the IRBP promoter activity in vitro, but their role in the control of IRBP gene expression in vivo is uncertain. Likely candidates for transcription factors that activate IRBP gene transcription and control its tissue-specific expression in adult photoreceptors are Crx, Nrl, and Otx2. These photoreceptor transcription factors do not work independently in the regulation of IRBP gene expression because the proteins pairs of Crx/Nrl and Otx2/Nrl work synergistically in transactivation assays [88, 95]. Interactions between these transcription factors may be different in rods versus cones. Crx transactivates the IRBP promoter in vitro. When mice carrying a transgene that consists of the IRBP promoter with a mutation in the CRX element upstream of a reporter gene, reporter gene expression is not found in photoreceptor cells of any of the transgenic mouse lines, indicating that the CRX element is necessary for IRBP expression in vivo [86]. Expression patterns of IRBP and the Crx element during retina development in the mouse are very similar to each other. The Crx message is first detected at E10.5 by rtPCR in mouse embryonic eyes [36], while IRBP is detected by RPA at E11.5 [35]. In the mouse, E12.5 is the time of cone cell genesis [33]. CRx protein is found in the neural developing retina at E12.5, and it is highly expressed in the adult [73, 74]. Surprisingly, however, IRBP expression levels appear unchanged in Crx null mice compared to wildtype mice [97], suggesting that Crx expression is not required for IRBP expression. Suppression of Crx by antisense treatment results in loss of promoter activity for p70, suggesting that while Crx is not absolutely requisite (Otx2 may be substituting), it may be the endogenous factor under normal (e.g., not knockout) situations [100]. Otx2 has an implied function in the activation of the IRBP gene; however, Otx2 expression in the adult is weak, suggesting that Otx2 is not the primary activator of IRBP gene expression in the adult. But, Crx expression is robust in adult photoreceptor cells. Crx and Otx2 are both members of the Otx gene family, which contains the

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paired class of homeodomain within their proteins. Homeodomain-containing proteins bind to DNA as a monomer, different homeodomain proteins can bind to the same DNA sequences, and other mechanisms are needed to achieve functional specificity for specific homeodomain transcription factors [117]. Perhaps there is a redundancy of function between Crx and Otx2. In the Crx null mice in which IRBP expression is basically normal, adequate amounts of Otx2 protein may be present and sufficient to maintain IRBP expression levels in the adult. Another transcription factor that may be required for IRBP gene expression is Rx/ rax, which is known to transactivate the IRBP promoter in vitro. During development, Rx is located at the correct time and place to activate IRBP gene expression in early retinal development. However, later in development, Rx expression in the retina becomes very low at E13.5 [118], whereas there is robust IRBP expression at this age. This indicates that the Rx/rax gene expression does not have a role in IRBP gene expression in the adult. Clearly, there are transcription factors that are required for the correct spatial and temporal expression of the IRBP gene that are yet to be identified. The retina is an excellent model for the study of brain development because it is part of the central nervous system, is easily accessible, and has a layered organization. Unlike many transcription factors found in the brain, retina-specific proteins are not required for viability and fertility, so null mutations do not have an adverse affect on the overall health of the animal. The mechanisms of cell fate determination and the gene families expressed by the progenitor cells are shared by the retina, cortex, and cerebellum. ACKNOWLEDGMENTS This work was supported by NIH EY11726 (D.E.B.), USU-CO70VY (D.E.B.), EY12514 (J.B.), R01EY014026 (J.B.), R01EY016470, R03EY013986, R24EY017045, and P30EY006360; the Foundation Fighting Blindness; Fight for Sight; Research to Prevent Blindness Inc.; and Knights Templar of Georgia. REFERENCES 1. Crouch RK, Hazard ES, Lind T, Wiggert B, Chader G, Corson DW. Interphotoreceptor retinoid-binding protein and alpha-tocopherol preserve the isomeric and oxidation state of retinol. Photochem Photobiol 1992;56(2):251–255. 2. Pfeffer B, Wiggert B, Lee L, Zonnenberg B, Newsome D, Chader G. The presence of a soluble interphotoreceptor retinol-binding protein (IRBP) in the retinal interphotoreceptor space. J Cell Physiol 1983;117(3):333–341. 3. Wiggert B, Lee L, Rodrigues M, Hess H, Redmond TM, Chader GJ. Immunochemical distribution of interphotoreceptor retinoid-binding protein in selected species. Invest Ophthalmol Vis Sci 1986;27(7):1041–1049. 4. Rodrigues MM, Hackett J, Gaskins R, Wiggert B, Lee L, Redmond M, et al. Interphotoreceptor retinoid-binding protein in retinal rod cells and pineal gland. Invest Ophthalmol Vis Sci 1986;27(5):844–850. 5. Chen Y, Saari JC, Noy N. Interactions of all-trans-retinol and long-chain fatty acids with interphotoreceptor retinoid-binding protein. Biochemistry 1993;32(42):11311–11318. 6. Chen Y, Houghton LA, Brenna JT, Noy N. Docosahexaenoic acid modulates the interactions of the interphotoreceptor retinoid-binding protein with 11-cis-retinal. J Biol Chem 1996;271(34):20507–20515.

116

Borst et al.

7. Fong SL, Liou GI, Landers RA, Alvarez RA, Bridges CD. Purification and characterization of a retinol-binding glycoprotein synthesized and secreted by bovine neural retina. J Biol Chem 1984;259(10):6534–6542. 8. Lai YL, Wiggert B, Liu YP, Chader GJ. Interphotoreceptor retinol-binding proteins: possible transport vehicles between compartments of the retina. Nature 1982;298(5877):848–849. 9. Saari JC, Teller DC, Crabb JW, Bredberg L. Properties of an interphotoreceptor retinoidbinding protein from bovine retina. J Biol Chem 1985;260(1):195–201. 10. Nickerson JM, Li GR, Lin ZY, Takizawa N, Si JS, Gross EA. Structure-function relationships in the four repeats of human interphotoreceptor retinoid-binding protein (IRBP). Mol Vis 1998;4:33. 11. Lin ZY, Li GR, Takizawa N, Si JS, Gross EA, Richardson K, et al. Structure-function relationships in interphotoreceptor retinoid-binding protein (IRBP). Mol Vis 1997;3:17. 12. Lin ZY, Si JS, Nickerson JM. Biochemical and biophysical properties of recombinant human interphotoreceptor retinoid binding protein. Invest Ophthalmol Vis Sci 1994;35(10):3599–3612. 13. Baer CA, Retief JD, Van Niel E, Braiman MS, Gonzalez-Fernandez F. Soluble expression in E. coli of a functional interphotoreceptor retinoid-binding protein module fused to thioredoxin: correlation of vitamin A binding regions with conserved domains of C-terminal processing proteases. Exp Eye Res 1998;66(2):249–262. 14. Baer CA, Kittredge KL, Klinger AL, Briercheck DM, Braiman MS, Gonzalez-Fernandez F. Expression and characterization of the fourth repeat of Xenopus interphotoreceptor retinoidbinding protein in E. coli. Curr Eye Res 1994;13(6):391–400. 15. Gonzalez-Fernandez F, Kittredge KL, Rayborn ME, Hollyfield JG, Landers RA, Saha M, et al. Interphotoreceptor retinoid-binding protein (IRBP), a major 124 kDa glycoprotein in the interphotoreceptor matrix of Xenopus laevis. Characterization, molecular cloning and biosynthesis. J Cell Sci 1993;105(Pt 1):7–21. 16. Hessler RB, Baer CA, Bukelman A, Kittredge KL, Gonzalez-Fernandez F. Interphotoreceptor retinoid-binding protein (IRBP): expression in the adult and developing Xenopus retina. J Comp Neurol 1996;367(3):329–341. 17. Liou GI, Fei Y, Peachey NS, Matragoon S, Wei S, Blaner WS, et al. Early onset photoreceptor abnormalities induced by targeted disruption of the interphotoreceptor retinoid-binding protein gene. J Neurosci 1998;18(12):4511–4520. 18. Ripps H, Peachey NS, Xu X, Nozell SE, Smith SB, Liou GI. The rhodopsin cycle is preserved in IRBP “knockout” mice despite abnormalities in retinal structure and function. Vis Neurosci 2000;17(1):97–105. 19. Donmoyer CM, Ciavatta VT, Liou GI, Nickerson JM. Microarray analysis of the neonatal Irbp knockout mouse. Invest Ophthalmol Vis Sci 2006;47:E-Abstract 4895. 20. Chader GJ. Retinal degenerations of hereditary, viral and autoimmune origins: studies on opsin and IRBP. Prog Retin Eye Res 1994;13(1):65–99. 21. Redmond TM, Sanui H, Nickerson JM, Borst DE, Wiggert B, Kuwabara T, et al. Cyanogen bromide fragments of bovine interphotoreceptor retinoid-binding protein induce experimental autoimmune uveoretinitis in Lewis rats. Curr Eye Res 1988;7(4):375–385. 22. Redmond TM, Si JS, Barrett DJ, Borst DE, Rainier S, Kotake S, et al. Synthesis of an immunopathogenic fusion protein derived from a bovine interphotoreceptor retinoid-binding protein cDNA clone. Gene 1989;80(1):109–118. 23. Kotake S, Wiggert B, Redmond TM, Borst DE, Nickerson JM, Margalit H, et al. Repeated determinants within the retinal interphotoreceptor retinoid-binding protein (IRBP): immunological properties of the repeats of an immunodominant determinant. Cell Immunol 1990;126(2):331–342. 24. Xu H, Wawrousek EF, Redmond TM, Nickerson JM, Wiggert B, Chan CC, et al. Transgenic expression of an immunologically privileged retinal antigen extraocularly enhances self

IRBP Molecular Biology

25.

26.

27. 28. 29. 30. 31. 32. 33. 34.

35.

36.

37.

38. 39.

40.

41. 42. 43.

117

tolerance and abrogates susceptibility to autoimmune uveitis. Eur J Immunol 2000;30(1): 272–278. Avichezer D, Liou GI, Chan CC, Lewis GM, Wiggert B, Donoso LA, et al. Interphotoreceptor retinoid-binding protein (IRBP)-deficient C57BL/6 mice have enhanced immunological and immunopathogenic responses to IRBP and an altered recognition of IRBP epitopes. J Autoimmun 2003;21(3):185–194. Grajewski RS, Silver PB, Agarwal RK, Su SB, Chan CC, Liou GI, et al. Endogenous IRBP can be dispensable for generation of natural CD4+CD25+ regulatory T cells that protect from IRBP-induced retinal autoimmunity. J Exp Med 2006;203(4):851–856. Turner DL, Cepko CL. A common progenitor for neurons and glia persists in rat retina late in development. Nature 1987;328(6126):131–136. Holt CE, Bertsch TW, Ellis HM, Harris WA. Cellular determination in the Xenopus retina is independent of lineage and birth date. Neuron 1988;1(1):15–26. Wetts R, Fraser SE. Multipotent precursors can give rise to all major cell types of the frog retina. Science 1988;239(4844):1142–1145. Turner DL, Snyder EY, Cepko CL. Lineage-independent determination of cell type in the embryonic mouse retina. Neuron 1990;4(6):833–845. Young RW. Cell differentiation in the retina of the mouse. Anat Rec 1985;212:199–205. Rapaport DH, Wong LL, Wood ED, Yasumura D, LaVail MM. Timing and topography of cell genesis in the rat retina. J Comp Neurol 2004;474(2):304–324. Carter-Dawson LD, LaVail MM. Rods and cones in the mouse retina. II. Autoradiographic analysis of cell generation using tritiated thymidine. J Comp Neurol 1979;188(2):263–272. Carter-Dawson L, Alvarez RA, Fong SL, Liou GI, Sperling HG, Bridges CD. Rhodopsin, 11-cis vitamin A, and interstitial retinol-binding protein (IRBP) during retinal development in normal and rd mutant mice. Dev Biol 1986;116(2):431–438. Liou GI, Wang M, Matragoon S. Timing of interphotoreceptor retinoid-binding protein (IRBP) gene expression and hypomethylation in developing mouse retina. Dev Biol 1994;161(2):345–356. Bibb LC, Holt JK, Tarttelin EE, Hodges MD, Gregory-Evans K, Rutherford A, et al. Temporal and spatial expression patterns of the CRX transcription factor and its downstream targets. Critical differences during human and mouse eye development. Hum Mol Genet 2001;10(15):1571–1579. Farr EA, Borst DE, Chader GJ, Bradley DJ. Developmental regulation of the mouse interphotoreceptor retinol binding protein gene in the fetal and post-natal eye. Invest Ophthalmol Vis Sci 1994;34:1727, Abstract 2199. O’Brien KM, Schulte D, Hendrickson AE. Expression of photoreceptor-associated molecules during human fetal eye development. Mol Vis 2003;9:401–409. Lee TC, Almeida D, Claros N, Abramson DH, Cobrinik D. Cell cycle-specific and cell type-specific expression of rb in the developing human retina. Invest Ophthalmol Vis Sci 2006;47(12):5590–5598. Kutty G, Duncan T, Nickerson JM, Si JS, Van Veen T, Chader GJ, et al. Light deprivation profoundly affects gene expression of interphotoreceptor retinoid-binding protein in the mouse eye. Exp Eye Res 1994;58(1):65–75. Katz ML, Gao CL, Stientjes HJ. Regulation of the interphotoreceptor retinoid-binding protein content of the retina by vitamin A. Exp Eye Res 1993;57(4):393–401. Fong SL, Bridges CD. Internal quadruplication in the structure of human interstitial retinolbinding protein deduced from its cloned cDNA. J Biol Chem 1988;263(30):15330–15334. Borst DE, Redmond TM, Elser JE, Gonda MA, Wiggert B, Chader GJ, et al. Interphotoreceptor retinoid-binding protein. Gene characterization, protein repeat structure, and its evolution. J Biol Chem 1989;264(2):1115–1123.

118

Borst et al.

44. Barrett DJ, Redmond TM, Wiggert B, Oprian DD, Chader GJ, Nickerson JM. cDNA clones encoding bovine interphotoreceptor retinoid binding protein. Biochem Biophys Res Commun 1985;131(3):1086–1093. 45. Liou GI, Ma DP, Yang YW, Geng L, Zhu C, Baehr W. Human interstitial retinoid-binding protein. Gene structure and primary structure. J Biol Chem 1989;264(14):8200–8206. 46. Inouye LN, Albini A, Chader GJ, Redmond TM, Nickerson JM. mRNA for interphotoreceptor retinoid-binding protein (IRBP): distribution and size diversity in vertebrate species. Exp Eye Res 1989;49(2):171–180. 47. Liou GI, Geng L, Baehr W. Interphotoreceptor retinoid-binding protein: biochemistry and molecular biology. Prog Clin Biol Res 1991;362:115–137. 48. Stenkamp DL, Calderwood JL, Van Niel EE, Daniels LM, Gonzalez-Fernandez F. The interphotoreceptor retinoid-binding protein (IRBP) of the chicken (Gallus gallus domesticus). Mol Vis 2005;11:833–845. 49. Rajendran RR, Van Niel EE, Stenkamp DL, Cunningham LL, Raymond PA, GonzalezFernandez F. Zebrafish interphotoreceptor retinoid-binding protein: differential circadian expression among cone subtypes. J Exp Biol 1996;199(Pt 12):2775–2787. 50. Stenkamp DL, Cunningham LL, Raymond PA, Gonzalez-Fernandez F. Novel expression pattern of interphotoreceptor retinoid-binding protein (IRBP) in the adult and developing zebrafish retina and RPE. Mol Vis 1998;4:26. 51. Nickerson JM, Frey RA, Ciavatta VT, Stenkamp DL. Interphotoreceptor retinoid-binding protein gene structure in tetrapods and teleost fish. Mol Vis 2006;12:1565–1585. 52. Nickerson JM, Borst DE, Redmond TM, Si JS, Toffenetti J, Chader GJ. The molecular biology of IRBP: application to problems of uveitis, protein chemistry, and evolution. Prog Clin Biol Res 1991;362:139–161. 53. van Veen T, Katial A, Shinohara T, Barrett DJ, Wiggert B, Chader GJ, et al. Retinal photoreceptor neurons and pinealocytes accumulate mRNA for interphotoreceptor retinoid-binding protein (IRBP). FEBS Lett 1986;208(1):133–137. 54. Porrello K, Bhat SP, Bok D. Detection of interphotoreceptor retinoid binding protein (IRBP) mRNA in human and cone-dominant squirrel retinas by in situ hybridization. J Histochem Cytochem 1991;39(2):171–176. 55. Egwuagu CE, Charukamnoetkanok P, Gery I. Thymic expression of autoantigens correlates with resistance to autoimmune disease. J Immunol 1997;159(7):3109–3112. 56. Fong SL, Balakier H, Canton M, Bridges CD, Gallie B. Retinoid-binding proteins in retinoblastoma tumors. Cancer Res 1988;48(5):1124–1128. 57. Fong SL, Fong WB, Morris TA, Kedzie KM, Bridges CD. Characterization and comparative structural features of the gene for human interstitial retinol-binding protein. J Biol Chem 1990;265(7):3648–3653. 58. Shuler RK, Jr., Gross E, He WY, Liou GI, Nickerson JM. Sequence analysis of the mouse IRBP gene and cDNA. Curr Eye Res 2002;24(5):354–367. 59. Borst DE, Boatright JH, Si JS, Stodulkova E, Remaley N, Pallansch LA, et al. Structural characterization and comparison of promoter activity of mouse and bovine interphotoreceptor retinoid-binding protein (IRBP) gene 5′ flanking regions in WERI, Y79, chick retina cells, and transgenic mice. Curr Eye Res 2001;23(1):20–32. 60. Javahery R, Khachi A, Lo K, Zenzie-Gregory B, Smale ST. DNA sequence requirements for transcriptional initiator activity in mammalian cells. Mol Cell Biol 1994;14(1):116–127. 61. Gonzalez-Fernandez F. Evolution of the visual cycle: the role of retinoid-binding proteins. J Endocrinol 2002;175(1):75–88. 62. Hoegg S, Brinkmann H, Taylor JS, Meyer A. Phylogenetic timing of the fish-specific genome duplication correlates with the diversification of teleost fish. J Mol Evol 2004;59(2):190–203.

IRBP Molecular Biology

119

63. Force A, Lynch M, Pickett FB, Amores A, Yan YL, Postlethwait J. Preservation of duplicate genes by complementary, degenerative mutations. Genetics 1999;151(4):1531–1545. 64. Postlethwait J, Amores A, Cresko W, Singer A, Yan YL. Subfunction partitioning, the teleost radiation and the annotation of the human genome. Trends Genet 2004;20(10):481–490. 65. Ohno S. Evolution by gene duplication. New York: Springer-Verlag; 1970. 66. Kadonaga JT. Regulation of RNA polymerase II transcription by sequence-specific DNA binding factors. Cell 2004;116(2):247–257. 67. Wray GA, Hahn MW, Abouheif E, Balhoff JP, Pizer M, Rockman MV, et al. The evolution of transcriptional regulation in eukaryotes. Mol Biol Evol 2003;20(9):1377–1419. 68. Kingston RE, Green MR. Modeling eukaryotic transcriptional activation. Curr Biol 1994;4(4):325–332. 69. Pabo CO, Sauer RT. Transcription factors: structural families and principles of DNA recognition. Annu Rev Biochem 1992;61:1053–1095. 70. Boatright JH, Knox BE, Jones KM, Stodulkova E, Nguyen HT, Padove SA, et al. Evidence of a tissue-restricting DNA regulatory element in the mouse IRBP promoter. FEBS Lett 2001;504(1–2):27–30. 71. Boatright JH, Borst DE, Peoples JW, Bruno J, Edwards CL, Si JS, et al. A major cis activator of the IRBP gene contains CRX-binding and Ret-1/PCE-I elements. Mol Vis 1997;3:15. 72. Werner M, Madreperla S, Lieberman P, Adler R. Expression of transfected genes by differentiated, postmitotic neurons and photoreceptors in primary cell cultures. J Neurosci Res 1990;25(1):50–57. 73. Furukawa T, Morrow EM, Cepko CL. Crx, a novel otx-like homeobox gene, shows photoreceptorspecific expression and regulates photoreceptor differentiation. Cell 1997;91(4):531–541. 74. Chen S, Wang QL, Nie Z, Sun H, Lennon G, Copeland NG, et al. Crx, a novel Otx-like paired-homeodomain protein, binds to and transactivates photoreceptor cell-specific genes. Neuron 1997;19(5):1017–1030. 75. Gumucio DL, Heilstedt-Williamson H, Gray TA, Tarle SA, Shelton DA, Tagle DA, et al. Phylogenetic footprinting reveals a nuclear protein which binds to silencer sequences in the human gamma and epsilon globin genes. Mol Cell Biol 1992;12(11):4919–4929. 76. Liou GI, Matragoon S, Overbeek PA, Fei Y. Identification of a retina-specific footprint within the retina-specific regulatory region of the human interphotoreceptor retinoid-binding protein gene. Biochem Biophys Res Commun 1994;203(3):1875–1881. 77. Boatright JH, Nickerson JM, Borst DE. Site-specific DNA hypomethylation permits expression of the IRBP gene. Brain Res 2000;887(1):211–221. 78. Freund CL, Gregory-Evans CY, Furukawa T, Papaioannou M, Looser J, Ploder L, et al. Cone-rod dystrophy due to mutations in a novel photoreceptor-specific homeobox gene (CRX) essential for maintenance of the photoreceptor. Cell 1997;91(4):543–553. 79. Swain PK, Chen S, Wang QL, Affatigato LM, Coats CL, Brady KD, et al. Mutations in the cone-rod homeobox gene are associated with the cone-rod dystrophy photoreceptor degeneration. Neuron 1997;19(6):1329–1336. 80. Borst DE, Si JS, Nickerson JM. Transient expression of IRBP promoter/CAT constructs in retinoblastoma cells. Invest Ophthalmol Vis Sci 1990;31:585, Abstract 2866. 81. Liou GI, Geng L, al-Ubaidi MR, Matragoon S, Hanten G, Baehr W, et al. Tissue-specific expression in transgenic mice directed by the 5′-flanking sequences of the human gene encoding interphotoreceptor retinoid-binding protein. J Biol Chem 1990;265(15):8373–8376. 82. Borst DE, Si JS, Nickerson JM. Functional comparison of the mouse IRBP promoter in Y79 cells and transgenic mice. Invest Ophthalmol Vis Sci 1992;33(4):1399, Abstract 3535. 83. Yokoyama T, Liou GI, Caldwell RB, Overbeek PA. Photoreceptor-specific activity of the human interphotoreceptor retinoid-binding protein (IRBP) promoter in transgenic mice. Exp Eye Res 1992;55(2):225–233.

120

Borst et al.

84. Liou GI, Matragoon S, Yang J, Geng L, Overbeek PA, Ma DP. Retina-specific expression from the IRBP promoter in transgenic mice is conferred by 212 bp of the 5′-flanking region. Biochem Biophys Res Commun 1991;181(1):159–165. 85. Bobola N, Hirsch E, Albini A, Altruda F, Noonan D, Ravazzolo R. A single cis-acting element in a short promoter segment of the gene encoding the interphotoreceptor retinoidbinding protein confers tissue-specific expression. J Biol Chem 1995;270(3):1289–1294. 86. Fei Y, Matragoon S, Smith SB, Overbeek PA, Chen S, Zack DJ, et al. Functional dissection of the promoter of the interphotoreceptor retinoid-binding protein gene: the cone-rod-homeobox element is essential for photoreceptor-specific expression in vivo. J Biochem (Tokyo) 1999;125(6):1189–1199. 87. Boatright JH, Knox BE, Stodulkova E, Nguyen HT, Padove SA, Matheu EA, et al. Tissuerestricting DNA regulatory elements in the mouse IRBP promoter. invest Ophthalmol Vis Sci 2001;42(4):S353, Abstract 1910. 88. Otteson DC, Liu Y, Lai H, Wang C, Gray S, Jain MK, et al. Kruppel-like factor 15, a zincfinger transcriptional regulator, represses the rhodopsin and interphotoreceptor retinoidbinding protein promoters. Invest Ophthalmol Vis Sci 2004;45(8):2522–2530. 89. Otteson DC, Lai H, Liu Y, Zack DJ. Zinc-finger domains of the transcriptional repressor KLF15 bind multiple sites in rhodopsin and IRBP promoters including the CRS-1 and Grich repressor elements. BMC Mol Biol 2005;6:15. 90. Fong SL, Fong WB. Elements regulating the transcription of human interstitial retinoidbinding protein (IRBP) gene in cultured retinoblastoma cells. Curr Eye Res 1999;18(4): 283–291. 91. Kikuchi T, Raju K, Breitman ML, Shinohara T. The proximal promoter of the mouse arrestin gene directs gene expression in photoreceptor cells and contains an evolutionarily conserved retinal factor-binding site. Mol Cell Biol 1993;13(7):4400–4408. 92. Kimura A, Singh D, Wawrousek EF, Kikuchi M, Nakamura M, Shinohara T. Both PCE1/RX and OTX/CRX interactions are necessary for photoreceptor-specific gene expression. J Biol Chem 2000;275(2):1152–1160. 93. Furukauea T, Kozak CA, Cepko CL. rax, a novel paired type homeobox gene, shows expressions in anterior neural fold and developing retuia. Proc Nalt Acad Sci USA 1997;94(7): 3088–3093. 94. Mathers, PH, Grenberg A, Mahon KA, Jamrich M. The Rx homeobox gene is essential for nertekrate eye development. Nature 1997;387(6633):603–607. 95. Akimoto M, Cheng H, Zhu D, Brzezinski JA, Khanna R, Filippova E, et al. Targeting of GFP to newborn rods by Nrl promoter and temporal expression profiling of flow-sorted photoreceptors. Proc Natl Acad Sci U S A 2006;103(10):3890–3895. 96. Mears AJ, Kondo M, Swain PK, Takada Y, Bush RA, Saunders TL, et al. Nrl is required for rod photoreceptor development. Nat Genet 2001;29(4):447–452. 97. Peng GH, Chen S. Chromatin immunoprecipitation identifies photoreceptor transcription factor targets in mouse models of retinal degeneration: new findings and challenges. Vis Neurosci 2005;22(5):575–586. 98. DuBridge RB, Tang P, Hsia HC, Leong PM, Miller JH, Calos MP. Analysis of mutation in human cells by using an Epstein-Barr virus shuttle system. Mol Cell Biol 1987;7(1):379–387. 99. Furukawa T, Morrow EM, Li T, Davis FC, Cepko CL. Retinopathy and attenuated circadian entrainment in Crx-deficient mice. Nat Genet 1999;23(4):466–470. 100. Livesey FJ, Furukawa T, Steffen MA, Church GM, Cepko CL. Microarray analysis of the transcriptional network controlled by the photoreceptor homeobox gene Crx. Curr Biol 2000;10(6):301–310.

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121

101. Bobola N, Briata P, Ilengo C, Rosatto N, Craft C, Corte G, et al. OTX2 homeodomain protein binds a DNA element necessary for interphotoreceptor retinoid binding protein gene expression. Mech Dev 1999;82(1–2):165–169. 102. Boatright JH, Borst DE, Stodulkova E, Nickerson JM. Endogenous CRX expression and IRBP promoter activity in retinoblastoma cells. Brain Res 2001;916(1–2):136–142. 103. Baas D, Bumsted KM, Martinez JA, Vaccarino FM, Wikler KC, Barnstable CJ. The subcellular localization of Otx2 is cell-type specific and developmentally regulated in the mouse retina. Brain Res Mol Brain Res 2000;78(1–2):26–37. 104. Ang SL, Jin O, Rhinn M, Daigle N, Stevenson L, Rossant J. A targeted mouse Otx2 mutation leads to severe defects in gastrulation and formation of axial mesoderm and to deletion of rostral brain. Development 1996;122(1):243–252. 105. Nishida A, Furukawa A, Koike C, Tano Y, Aizawa S, Matsuo I, et al. Otx2 homeobox gene controls retinal photoreceptor cell fate and pineal gland development. Nat Neurosci 2003;6(12):1255–1263. 106. Courtois V, Chatelain G, Han ZY, Le Novere N, Brun G, Lamonerie T. New Otx2 mRNA isoforms expressed in the mouse brain. J Neurochem 2003;84(4):840–853. 107. Arranz V, Dreuillet C, Crisanti P, Tillit J, Kress M, Ernoult-Lange M. The zinc finger transcription factor, MOK2, negatively modulates expression of the interphotoreceptor retinoid-binding protein gene, IRBP. J Biol Chem 2001;276(15):11963–11969. 108. Livne-Bar I, Pacal M, Cheung MC, Hankin M, Trogadis J, Chen D, et al. Chx10 is required to block photoreceptor differentiation but is dispensable for progenitor proliferation in the postnatal retina. Proc Natl Acad Sci U S A 2006;103(13):4988–4993. 109. Liu IS, Chen JD, Ploder L, Vidgen D, van der Kooy D, Kalnins VI, et al. Developmental expression of a novel murine homeobox gene (Chx10): evidence for roles in determination of the neuroretina and inner nuclear layer. Neuron 1994;13(2):377–393. 110. Dorval KM, Bobechko BP, Ahmad KF, Bremner R. Transcriptional activity of the pairedlike homeodomain proteins Chx10 and VSX1. J Biol Chem 2005;280(11):10100–10108. 111. Rowan S, Cepko CL. Genetic analysis of the homeodomain transcription factor Chx10 in the retina using a novel multifunctional BAC transgenic mouse reporter. Dev Biol 2004;271(2):388–402. 112. Rutherford AD, Dhomen N, Smith HK, Sowden JC. Delayed expression of the Crx gene and photoreceptor development in the Chx10-deficient retina. Invest Ophthalmol Vis Sci 2004;45(2):375–384. 113. Dorval KM, Bobechko BP, Fujieda H, Chen S, Zack DJ, Bremner R. Chx10 targets a subset of photoreceptor genes. J Biol Chem 2006;281(2):744–751. 114. Funk WD, Wright WE. Cyclic amplification and selection of targets for multicomponent complexes: myogenin interacts with factors recognizing binding sites for basic helix-loophelix, nuclear factor 1, myocyte-specific enhancer-binding factor 2, and COMP1 factor. Proc Natl Acad Sci U S A 1992;89(20):9484–9488. 115. Knoepfler PS, Bergstrom DA, Uetsuki T, Dac-Korytko I, Sun YH, Wright WE, et al. A conserved motif N-terminal to the DNA-binding domains of myogenic bHLH transcription factors mediates cooperative DNA binding with pbx-Meis1/Prep1. Nucleic Acids Res 1999;27(18):3752–3761. 116. Ferretti E, Villaescusa JC, Di Rosa P, Fernandez-Diaz LC, Longobardi E, Mazzieri R, et al. Hypomorphic mutation of the TALE gene Prep1 (pKnox1) causes a major reduction of Pbx and Meis proteins and a pleiotropic embryonic phenotype. Mol Cell Biol 2006;26(15):5650–562. 117. Hisa T, Spence SE, Rachel RA, Fujita M, Nakamura T, Ward JM, et al. Hematopoietic, angiogenic and eye defects in Meis1 mutant animals. EMBO J 2004;23(2):450–459.

122

Borst et al.

118. Zhang X, Friedman A, Heaney S, Purcell P, Maas RL. Meis homeoproteins directly regulate Pax6 during vertebrate lens morphogenesis. Genes Dev 2002;16(16):2097–2107. 119. Wilson D, Sheng G, Lecuit T, Dostatni N, Desplan C. Cooperative dimerization of paired class homeo domains on DNA. Genes Dev 1993;7(11):2120–2134. 120. Mathers PH, Grinberg A, Mahon KA, Jamrich M. The Rx homeobox gene is essential for vertebrate eye development. Nature 1997;387(6633):603–607. 121. Nickerson JM, Frey RA, Ciavatta VT, Stenkamp DL. Interphotoreceptor retinoid-binding protein gene structure in tetrapods and teleost fish. Mol Vis 2006;12:1565–1585.

6 Regulation of Photoresponses by Phosphorylation Alecia K. Gross, Qiong Wang, and Theodore G. Wensel CONTENTS Introduction Inactivation of Photoactivated Rhodopsin by Rhodopsin Kinase Cone-Specific Kinase, GRK7 Protein Kinase C cAMP-Dependent Protein Kinase, PKA Cyclin-Dependent Kinase, CDK5 Tyrosine Kinases Mitogen-Activated Protein Kinase and Calmodulin-Dependent Protein Kinase II Protein Phosphatases Conclusion References

INTRODUCTION In common with virtually all signaling pathways in biology, phototransduction in rod and cone photoreceptors of the vertebrate retina is regulated by protein phosphorylation. Most of the attention of researchers has focused on rhodopsin kinase (RK), an enzyme unique to photoreceptors that is the essential first step for normal photoresponse recovery kinetics (Fig. 1). In addition to this important enzyme, myriad other protein kinases are expressed in photoreceptors, and their functions are much less clear. Along with protein kinases, the activity of protein phosphatases determines the dynamics of protein phosphorylation, and therefore activity, and these have been the subject of many previous and ongoing studies as well. Thus, phosphorylation of photoreceptor proteins, its regulation, and its functional consequences for photoresponses have been and remain active areas of research. Previous reviews provide further information on specific topics [1–15].

From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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Fig. 1. The role of phosphorylation by rhodopsin kinase in the inactivation of photoexcited rhodopsin, metarhodopsin II (MII or R*). In the dark-adapted rod, rhodopsin is not a good substrate for rhodopsin kinase (RK), so on photoactivation and conversion to R* it catalyzes rapid guanosine diphosphate–guanosine triphosphate (GDP-GTP) exchange on the α-subunit of the G protein transducin, Gαt. Activated Gαt-GTP activates cyclic guanosine monophosphate (cGMP) phosphodiesterase (PDE), leading to closure of cGMP-gated cation channels and plasma membrane hyperpolarization. R* binds to and activates rhodopsin kinase, which uses adenosine triphosphate (ATP) to add multiple phosphates to the carboxyl-terminal region of R*, inducing a state that binds with high affinity to arrestin. Arrestin binding to phosphorylated rhodopsin effectively quenches the activity of R* until decay of metarhodopsin II and regeneration of rhodopsin lead to dephosphorylation by protein phosphatase 2A (PP2A). Although the decay of metarhodopsin II and regeneration eventually lead to inactivation of the phototransduction cascade in the absence of rhodopsin kinase or its target sites, this process is much too slow for normal recovery kinetics. The presence of multiple phosphorylation sites enhances the reproducibility of inactivation kinetics by decreasing the variation in the amount of time each R* remains active.

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Fig. 2. Locations of domains and functional sites within the primary structure of rhodopsin kinase. Diamonds represent phosphorylation sites, with the numbers indicating sequence positions. The RGS domain is homologous to the catalytic domains of RGS proteins but is not known to demonstrate GAP (guanosine triphosphatase accelerating protein) activity. The kinase domain is homologous to other serine/threonine protein kinases. At the carboxyl terminus, the CaaX-box sequence CSVS is subject to posttranslational modification in which the last three amino acid residues are proteolytically cleaved, the cysteine is farnesylated, and the terminal carboxylate is converted to a methyl ester.

INACTIVATION OF PHOTOACTIVATED RHODOPSIN BY RHODOPSIN KINASE Inactivation of photoexcited rhodopsin (metarhodopsin II, MII, or R*) is initiated via its enzymatic phosphorylation by rhodopsin kinase (RK, Fig. 2). The activity of RK was first described 35 years ago [16–19] when isolated rod outer segments (ROSs) were incubated with γ-32P adenosine triphosphatase (ATP) in the presence of light, causing 32P to be incorporated into MII. This light-dependent process was shown to occur in vivo [20], and later phosphorylated rhodopsin was shown to be necessary for signal attenuation in bovine rod outer segment preparations [2, 3] and subsequently in vivo [21]. After over a decade of attempts, RK was purified, stabilized, and characterized [22]. Since then, experiments from many laboratories have shed light on both the properties of the enzyme and its importance in the visual transduction process. Rhodopsin kinase, also known as G protein-coupled receptor kinase 1 (GRK1), was the first member discovered of the GRK family of Ser/Thr kinases specific for seventransmembrane G protein-coupled receptors [23]. It is responsible for phosphorylating MII on its C-terminus (see structures in Fig. 3) with an upper limit of nine phosphorylation sites per MII [24]. This phosphorylation allows the binding of arrestin to occur, effectively quenching the signaling pathway by no longer allowing the G-protein transducin (Gt) to interact with MII [25]. Depending on reaction conditions, addition of phosphates to the C-terminus of rhodopsin can either decrease [25, 26] or fully attenuate [27] its interaction with transducin. Rhodopsin kinase is a single-polypeptide chain enzyme with a molecular weight of 62–64 kDa [28] and is posttranslationally modified: it is farnesylated at the C-terminus consensus sequence for isoprenylation, CaaX, which is followed by limited proteolysis of aaX and subsequent methyl esterification of the isoprenylated Cys [29, 30] (Fig. 2). Without these modifications, the activity of RK is approximately four-fold lower, suggesting that this hydrophobic modification is important for targeting RK to disk membranes and conferring full enzymatic activity toward MII. RK is autophosphorylated on serine residues, and the reaction is unaffected by the presence of bleached rhodopsin [11–13]. This autophosphorylation does not alter the rate of rhodopsin phosphorylation.

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Fig. 3. Models of structures of the C-terminus (residues 330–348) of rhodopsin (left) from x-ray crystallography (PDB file 1U19) of the dark state of rhodopsin [131] and (right) from nuclear magnetic resonance (NMR) (PDB file 1NZS) of the peptide chemically phosphorylated at the seven labeled serine and threonine residues and bound to arrestin [132]. These structures suggest that photoactivation, phosphorylation, or arrestin binding may involve dramatic differences in the conformation of this region.

The kinetic parameters of the purified enzyme for MII are as follows: Km= 4 µM (rhodopsin), Km = 2 µM (ATP), and Vmax = 700 nmol-min−1-mg−1, corresponding to a turnover number, kcat = 0.8 s −1 [22]. However, in vivo, rhodopsin is inactivated with a time constant of 80 ms or less [31]. Proof that RK binding to MII is required for normal inactivation of MII in rods first came from recordings of photocurrent responses of single mouse rods expressing a C-terminal truncation mutant of rhodopsin [21] and subsequently from responses of mouse rods with a null mutation of RK [32]. Mutations in RK cause defects in the kinetics of deactivation and an increase in the amplitude of the light response. While RK can phosphorylate many serine and threonine residues on the C-terminus of rhodopsin, only a few (Ser 334, Ser338, and Ser343) have been identified biochemically as major phosphorylation sites in intact retinas [33, 34]. Based on experiments using rapid quench followed by mass spectrometry, it was reported that in mouse retinas the sites closest to the C-terminus of rhodopsin are the first to be phosphorylated; Ser343 is phosphorylated most rapidly, followed by Ser338. Ser334 is phosphorylated after a delay of more than 10 s [35]. In relating these experiments to electrophysiological results, it must be borne in mind that these experiments required supersaturating levels of light to obtain sufficient product for analysis, and therefore the kinetics may differ in the dim flash or single-photon regimes. The requirement for multiple biochemical steps to inactivate MII has been proposed to play an important role in reducing the variability in the lifetime of catalytically active

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Fig. 4. Structure of recoverin bound to an N-terminal peptide from rhodopsin kinase (RK), illustrating interactions that may help mediate Ca2+-dependent inhibition of this kinase. Structures (2I94) are based on nuclear magnetic resonance (NMR) [133] of recoverin in the presence of Ca2+, which is bound at the third and fourth EF hands, and a peptide, RK25, corresponding to the first 25 amino acid residues of rhodopsin kinase.

rhodopsin [36]. A recent electrophysiological study using rhodopsin transgenes encoding proteins with different numbers of phosphorylation sites [37] elegantly demonstrated the importance of the multiplicity of these sites in single-photon reproducibility. The activity of RK for isolated peptide substrates derived from the C-terminal region of rhodopsin is around 1,000-fold lower than for the full-length MII [38]. Peptide competition studies and alanine-scanning mutagenesis of rhodopsin have shown the interaction site of RK with rhodopsin to include cytoplasmic loops 1 through 3 [28, 39]. Early studies of rhodopsin phosphorylation have shown that at low light levels (sufficient to activate < 1% of the total rhodopsin pool), for every mole of activated rhodopsin several hundred moles of phosphate were added to the rhodopsin pool [17, 40]. A straightforward explanation of this phenomenon, known as high-gain phosphorylation, is that the nonspecific substrate, inactive rhodopsin, is being phosphorylated in a light-dependent manner in trans. While RK has been shown to exist in two states (an inactive and an active state using proteolytically digested rhodopsin from bovine rod outer segments and synthetic peptides [41]), trans-phosphorylation does not occur in heterologously expressed chimeras of rhodopsin [42], still leaving the underlying cause of transphosphorylation uncertain. Lack of functionally active RK in humans has been shown to cause blinding diseases: Oguchi disease, a form of congenital stationary night blindness, and retinitis pigmentosa. While Oguchi disease is characterized by profoundly slowed rod dark adaptation [43], patients with retinitis pigmentosa undergo progressive blindness [44]. The visual functions of a patient with an inactivating mutation in the RK gene have been thoroughly characterized by electrophysiological and psychophysical methods [45]. In mice, absence of RK leads to a light-dependent degeneration of the retina [32]. Rhodopsin kinase has been reported to be regulated by recoverin, a member of the neuronal calcium sensor (NCS) branch of the EF-hand superfamily [46, 47]. In vitro studies together with electrophysiological studies of recoverin knockout mice suggest that by inhibiting RK at high calcium levels (Fig. 4), recoverin prolongs the lifetime of MII [47], thereby allowing the accelerated inactivation of rhodopsin in response to lowered intracellular calcium under light conditions [48, 49].

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CONE-SPECIFIC KINASE, GRK7 While most work on phosphorylation of photoreceptors was performed in rod cells of rod-dominant mammals, GRK7, the enzyme involved in cone recovery, has been studied with the aid of cone-dominant animals such as ground squirrels and chipmunks, as well as the medaka Oryzias latipes [33, 34]. Interestingly, its expression is species specific: Immunocytochemical analysis has shown that GRK7 is expressed exclusively in pig and dog cones, whereas GRK1 is the only known photoreceptor GRK found in mice and rats [50]. In contrast, GRK7 and GRK1 are coexpressed in cones of humans and monkeys, suggesting coordination between the kinases in primates. Studies with carp cones suggest that their pigments are phosphorylated much more rapidly than rhodopsin is in rods from the same species [51]. Phosphorylation of M opsin by GRK7 was shown to reduce the ability of the photopigment to activate the G protein transducin in vitro using a filter-binding assay [52]. In addition, in retinal homogenates of both rod-dominated (pig) and cone-dominated (13-lined ground squirrel) animals, anti-ground squirrel GRK7 antibodies were shown to block light-dependent phosphorylation of M opsins to varying degrees, suggesting GRK7 is at least partially responsible for phosphorylation of M opsins.

PROTEIN KINASE C Substantial enzymatic activity and immunoreactivity (by immunoblotting, not by immunocytochemistry) of protein kinase C are found in photoreceptor outer segments [53–64], and specific isoforms identified include conventional (Ca2+-, phospholipid-, and diacyl glycerol/phorbol ester-dependent) isoform protein kinase Cα (PKCα) and novel (non-Ca2+-dependent) isoforms ε and θ [63, 65, 66]. There have been a number of reports concerning the identity of substrates for PKC in rod outer segments, but the significance and extent of their phosphorylation by PKC isoforms have remained uncertain in most cases. Rhodopsin was reported to be phosphorylated in its C-terminal tail region by PKC [56, 57, 61, 62, 64, 67, 68]. Indeed, in rod outer segments treated with phorbol esters, rhodopsin is the major phosphoprotein detected [62]. However, studies in RK knockout mice revealed no light-induced phosphorylation of rhodopsin residue Ser338 and Ser334, the major in vitro PKC phosphorylation sites, raising questions about the significance in vivo of PKC phosphorylation of rhodopsin. The photoreceptor G protein, transducin α or Gαt, has been reported to be a PKC substrate [15]. Addition of exogenous PKCβ1 led to Gαt phosphorylation in a manner dependent on the state of association with rhodopsin, guanine nucleotide, and Gβγt [69]. Both the α-catalytic subunit and the inhibitory γ-subunit of the cyclic guanosine monophosphate (cGMP) phosphodiesterase, PDE6, have been reported to be phosphorylated in vitro by purified PKC [1, 70, 71]. The physiological significance of this reaction remains unclear, and there do not seem to have been any reports of in situ phosphorylation of this effector enzyme by endogenous PKC. The photoreceptor guanylate cyclase (GC) has been reported to be enhanced by PKC [72], and expression of the rod outer segment-GC1 gene has been reported to be regulated by PKC [73].

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Fig. 5. Schematic diagram of regulation of RGS9-1 activity by protein kinase C α (PKCα) and Ca2+. In the dark with relatively high Ca2+ concentration and PKCα activity, RGS9-1 is phosphorylated. This phosphorylation reduces RGS9-1 GAP (guanosine triphosphatase [GTPase] accelerating protein) activity by inhibiting its interactions with the membrane anchor and GAP activator, R9AP, and thereby allows maximum sensitivity in the dark to allow detection of single photons. In the light, lowering of intracellular [Ca2+] reduces PKC activity so that RGS9-1 becomes dephosphorylated, presumably by protein phosphatase 2A (PP2A), allowing rapid shutoff of the cascade. ADP adenosine diphosphate, ATP adenosine triphosphate, GDP guanosine diphosphate, GTP guanosine triphosphate.

The photoreceptor GAP (guanosine triphosphatase [GTPase] accelerating protein) RGS9-1 is phosphorylated by PKCα (unpublished observations by Q.W. and T.G.W.) in a reaction that is controlled by Ca2+ in vitro and by light in vivo [66, 74]. This phosphorylation lowers the affinity of RGS9-1 for its membrane anchor, R9AP, whose binding dramatically increases the activity of RGS9-1 [75, 76]. An interesting anomaly of this reaction is that phorbol esters greatly enhance phosphorylation of rhodopsin in the dark but do not enhance PKC-mediated phosphorylation of RGS9-1 [74]. A model for regulation of RGS9-1 by PKC phosphorylation is proposed in Fig. 5. The physiological importance of these phosphorylation events is still under investigation. Electrophysiological experiments with individual salamander rods showed no effects of phorbol esters or a PKC inhibitor on light responses [77]; likewise, PKC inhibitors were found not to affect light responses in rods of the Tokay gecko [78]. Knockout mice are available for several PKC isoforms, but studies of their light responses have yet to be published.

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CYCLIC ADENOSINE MONOPHOSPHATE-DEPENDENT PROTEIN KINASE, PKA Cyclic nucleotides have long been known to stimulate phosphorylation of multiple proteins in rod outer segments through the action of protein kinase A (PKA) [79]. PKA has been reported to phosphorylate both pigment kinases, GRK1 and GRK7 [80], to regulate guanylate cyclase [81], and to phosphorylate PDE6 [82]. PKA has also been implicated in the activation of kinases that regulate the CNG (cyclic nucleotide-gated) channel [83], described in the section on mitogen-activated protein kinase (MAPK) and calmodulin-dependent protein kinase II (CamKII). In addition, PKA has been implicated in regulation of calcium currents at photoreceptor synapses by dopamine and adenosine receptors [84, 85]. It has been reported that PKA can phosphorylate RGS9-1 [86]; however, only PKC has been implicated clearly in light-dependent RGS9-1 phosphorylation in vivo (see preceding section). One of the major substrates for PKA in photoreceptors is phosducin [87–89]. In photoreceptors, phosducin is found primarily in the inner segment, where PKA maintains it in a phosphorylated state in the dark. In this state, it binds to 14-3-3 proteins in the inner segment [90]. Light leads to lowered activity of PKA toward phosducin, resulting in dephosphorylation, and in this state, phosducin binds tightly to the Gβ1γ1 subunit of Gt [91, 92]; a crystal structure of this complex has been solved [93, 94]. However, the subcellular localization of phosducin residing primarily in the inner segments as well as results from phosducin knockout mice [95–98] suggest that it does not directly regulate phototransduction in the outer segments. Rather, it seems to facilitate translocation of Gβ1γ1 in response to high levels of light. It has also been proposed to be involved in regulation of transcription [99, 100]. CYCLIN-DEPENDENT KINASE The cyclin-dependent kinase CDK5 and its regulatory subunit p35 are present in rod outer segments and can phosphorylate the inhibitory γ-subunit (PDE6γ) of the cGMP phosphodiesterase PDE6 [74, 101, 102]. A study of PDE6γ phosphorylated on Thr22, the site for phosphorylation by CDK5, revealed that the phosphoprotein has greatly diminished affinity for the activated form of the G protein Gαt-GTP and relatively little change in its inhibitory activity or affinity for the catalytic subunits of PDE6. PDE6γ apparently has to dissociate from the catalytic subunits of PDE6 to be phosphorylated efficiently, and it remains to be determined to what extent this occurs in vivo. TYROSINE KINASES Receptors for insulin and insulin-like growth factor 1 (IGF-1) have been reported to be present in rod outer segments [103–105]. The nonreceptor tyrosine kinases c-Src and p56lck are also present [106, 107], and c-Src was reported to bind to the α-subunit of the G protein (Gαt). Modulation of the photoreceptor cGMP-gated cation channel by IGF-1 and tyrosine phosphorylation/dephosphorylation has been reported [108, 109]. Light activation of rhodopsin has been reported to stimulate tyrosine phosphorylation of the insulin receptor β-subunit and activation of phosphoinositide-3 kinase [8, 110, 111]. The precise physiological functions of these tyrosine phosphorylation reactions will

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likely be the subject of further investigations. It has also been reported that fibroblast growth factor (FGF) effects on retinal survival involve direct effects on photoreceptor cells [112], and that the FGF receptor FGFR-1 is expressed in photoreceptors [113]. There have been reports of the neurotrophin receptors of the Trk family being present in photoreceptor cells [114–117], but the role of these receptors, known to be critical for proper development, in adult photoreceptors remains unclear. MITOGEN-ACTIVATED PROTEIN KINASE AND CALMODULIN-DEPENDENT PROTEIN KINASE II The MAPK Erk and CamKII have been reported to regulate cone cGMP-gated channels [118]. Erk, which is maximally active during subjective night, appears to enhance the affinity of the channel for cGMP, whereas CaMKII, which has maximum activity during the subjective day, has the opposite effect. CamKII has also been reported to phosphorylate phosducin (see the section on cAMP-dependent PKA) in the inner segment [90, 119]. PROTEIN PHOSPHATASES The phosphorylation state of any photoreceptor protein is determined by the balance of activities of protein kinases and protein phosphatases. Serine/threonine protein phosphatases 1, 2A, and 2C are all present in purified rod outer segments [120–123]. Protein phosphatase 2A (PP2A) is primarily responsible for dephosphorylation of phosphorylated opsin [124–126] and RGS9-1 [127] and has been reported to undergo translocation in response to light [128]. Tyrosine phosphatase SHP (Src-homology 2-domain phosphatase)-1 [129] and SHP-2 [130] are expressed in photoreceptors. SHP-1 deletion in mice caused severe retinal degeneration phenotype at 3 weeks of age [129]. SHP-2 was reported to be associated with Gαt in rod outer segments and to undergo light-activated phosphorylation [130]. Further investigation will be needed to determine the physiological substrates and roles of these tyrosine phosphatases. CONCLUSION The role of phosphorylation in regulating light responses has been studied fairly intensively for over two decades. The rich fund of information derived from these studies, especially in the area of rhodopsin phosphorylation, serves not only to help explain key elements of the recovery phases of light responses, but also to help us understand the roles of phosphorylation and arrestin binding in other G protein-mediated signaling pathways. Nonetheless, many intriguing questions remain. Most of these have to do with the roles of the many kinases other than RK that are found in photoreceptors, as well as questions about phosphatases and what sorts of regulation might control their activities. Important questions remain about rhodopsin phosphorylation as well. These include the kinetics of phosphorylation of specific target residues in MII not in response to oversaturating light intensities, but in the dim flash regime where responses are linear sums of single-photon responses. This range of intensities has so far not been accessible to biochemical techniques used to study phosphorylation kinetics in intact retinas. Additional

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remaining questions concern the mechanisms by which genetic defects in rhodopsin phosphorylation lead to retinal degeneration in some contexts and stationary night blindness in others. All of these will likely be the subjects of ongoing studies in the future. REFERENCES 1. Palczewski, K. (1997). GTP-binding-protein-coupled receptor kinases—two mechanistic models. Eur J Biochem 248, 261–269. 2. Baylor, D. A., Burns, M. E. (1998). Control of rhodopsin activity in vision. Eye 12, 521–5. 3. Penn, R. B., Pronin, A. N., Benovic, J. L. (2000). Regulation of G protein-coupled receptor kinases. Trends Cardiovasc Med 10, 81–89. 4. Maeda, T., Imanishi, Y., Palczewski, K. (2003). Rhodopsin phosphorylation: 30 years later. Prog Retin Eye Res 22, 417–434. 5. Dryja, T. P. (2000). Molecular genetics of Oguchi disease, fundus albipunctatus, and other forms of stationary night blindness: LVII Edward Jackson Memorial Lecture. Am J Ophthalmol 130, 547–563. 6. Hargrave, P. A., McDowell, J. H. (1992). Rhodopsin and phototransduction: a model system for G protein-linked receptors. Faseb J 6, 2323–2331. 7. Yamazaki, A., Moskvin, O., Yamazaki, R. K. (2002). Phosphorylation by cyclin-dependent protein kinase 5 of the regulatory subunit (Pgamma) of retinal cgmp phosphodiesterase (PDE6): its implications in phototransduction. Adv Exp Med Biol 514, 131–153. 8. Rajala, R. V., Anderson, R. E. (2003). Light regulation of the insulin receptor in the retina. Mol Neurobiol 28, 123–138. 9. Kawamura, S., Tachibanaki, S. (2002). S-modulin. Adv Exp Med Biol 514, 61–68. 10. Chen, C. K. (2002). Recoverin and rhodopsin kinase. Adv Exp Med Biol 514, 101–107. 11. Arshavsky, V. Y. (2002). Rhodopsin phosphorylation: from terminating single photon responses to photoreceptor dark adaptation. Trends Neurosci 25, 124–126. 12. Kawamura, S. (1999). Calcium-dependent regulation of rhodopsin phosphorylation. Novartis Found Symp 224, 208–218; discussion 218–224. 13. Hurley, J. B., Spencer, M., Niemi, G. A. (1998). Rhodopsin phosphorylation and its role in photoreceptor function. Vision Res 38, 1341–1352. 14. Palczewski, K., Saari, J. C. (1997). Activation and inactivation steps in the visual transduction pathway. Curr Opin Neurobiol 7, 500–504. 15. Zhao, X., Palczewski, K., Ohguro, H. (1995). Mechanism of rhodopsin phosphorylation. Biophys Chem 56, 183–188. 16. Frank, R. N., Cavanagh, H. D., Kenyon, K. R. (1973). Light-stimulated phosphorylation of bovine visual pigments by adenosine triphosphate. J Biol Chem. 248, 596–609. 17. Bownds, D., Dawes, J., Miller, J., Stahlman, M. (1972). Phosphorylation of frog photoreceptor membranes induced by light. Nature New Biology 237, 125–127. 18. Frank, R. N., Cavanagh, H. D., Kenyon, K. R. (1973). Light-stimulated phosphorylation of bovine visual pigments by adenosine triphosphate. J Biol Chem. 248, 596–609. 19. Kühn, H., Dreyer, W. J. (1972). Light dependent phosphorylation of rhodopsin by ATP. FEBS Lett. 20, 1–6. 20. Kuhn, H. (1974). Light-dependent phosphorylation of rhodopsin in living frogs. Nature 250, 588–590. 21. Chen, J., Makino, C. L., Peachey, N. S., Baylor, D. A., Simon, M. I. (1995). Mechanisms of rhodopsin inactivation in vivo as revealed by a COOH− terminal truncation mutant. Science 267, 374–377. 22. Palczewski, K., McDowell, J. H., Hargrave, P. A. (1988). Purification and characterization of rhodopsin kinase. J Biol Chem 263, 14067–14073.

Phosphorylation in Phototransduction

135

23. Palczewski, K., Benovic, J. L. (1991). G-protein coupled receptor kinases. Trends Biochem Sci 16, 387–391. 24. Wilden, U., Kühn, H. (1982). Light-dependent phosphorylation of rhodopsin: Number of phosphorylation sites. Biochemistry 21, 3014–3022. 25. Kühn, H., Hall, S. W., Wilden, U. (1984). Light-induced binding of 48-kDa protein to photoreceptor membranes is highly enhanced by phosphorylation of rhodopsin. FEBS Lett 176, 473–478. 26. Liebman, P. A., Pugh, E. N., Jr. (1982). Gain, speed and sensitivity of GTP binding vs PDE activation in visual excitation. Vision Res 22, 1475–1480. 27. Sitaramayya, A., Harkness, J., Parkes, J. H., Gonzalez-Oliva, C., Liebman, P. A. (1986). Kinetic studies suggest that light-activated cyclic GMP phosphodiesterase is a complex with G-protein subunits. Biochemistry 25, 651–656. 28. Kelleher, D. J., Johnson, G. L. (1990). Characterization of rhodopsin kinase purified from bovine rod outer segments. J Biol Chem 265, 2632–2639. 29. Inglese, J., Glickman, J. F., Lorenz, W., Caron, M. G., Lefkowitz, R. J. (1992). Isoprenylation of a protein kinase. Requirement of farnesylation/alpha-carboxyl methylation for full enzymatic activity of rhodopsin kinase. J Biol Chem 267, 1422–1425. 30. Inglese, J., Koch, W. J., Caron, M. G., Lefkowitz, R. J. (1992). Isoprenylation in regulation of signal transduction by G-protein-coupled receptor kinases. Nature 359, 147–150. 31. Krispel, C. M., Chen, D., Melling, N., Chen, Y. J., Martemyanov, K. A., Quillinan, N., Arshavsky, V. Y., Wensel, T. G., Chen, C. K., Burns, M. E. (2006). RGS expression ratelimits recovery of rod photoresponses. Neuron 51, 409–416. 32. Chen, C. K., Burns, M. E., Spencer, M., Niemi, G. A., Chen, J., Hurley, J. B., Baylor, D. A., Simon, M. I. (1999). Abnormal photoresponses and light-induced apoptosis in rods lacking rhodopsin kinase. Proc Natl Acad Sci U S A 96, 3718–3722. 33. Ohguro, H., Palczewski, K., Ericsson, L. H., Walsh, K. A., Johnson, R. S. (1993). Sequential phosphorylation of rhodopsin at multiple sites. Biochemistry 32, 5718–5724. 34. McDowell, J. H., Nawrocki, J. P., Hargrave, P. A. (1993). Phosphorylation sites in bovine rhodopsin Biochemistry 32, 4968–4974. 35. Kennedy, M. J., Lee, K. A., Niemi, G. A., Craven, K. B., Garwin, G. G., Saari, J. C., Hurley, J. B. (2001). Multiple phosphorylation of rhodopsin and the in vivo chemistry underlying rod photoreceptor dark adaptation. Neuron 31, 87–101. 36. Hamer, R. D., Nicholas, S. C., Tranchina, D., Liebman, P. A., Lamb, T. D. (2003). Multiple steps of phosphorylation of activated rhodopsin can account for the reproducibility of vertebrate rod single-photon responses. J Gen Physiol 122, 419–444. 37. Doan, T., Mendez, A., Detwiler, P. B., Chen, J., Rieke, F. (2006). Multiple phosphorylation sites confer reproducibility of the rod’s single-photon responses. Science 313, 530–533. 38. Palczewski, K., Arendt, A., McDowell, J. H., Hargrave, P. A. (1989). Substrate recognition determinants or rhodopsin kinase: studies with synthetic peptides, polyanions, and polycations. Biochemistry 28, 8764–8770. 39. Shi, W., Osawa, S., Dickerson, C. D., Weiss, E. R. (1995). Rhodopsin mutants discriminate sites important for the activation of rhodopsin kinase and Gt*. J Biol Chem 270, 2112–2119. 40. Kühn, H., Dreyer, W. J. (1972). Light dependent phosphorylation of rhodopsin by ATP. FEBS Lett 20, 1–6. 41. Palczewski, K., Buczylko, J., Kaplan, M. W., Polans, A. S., Crabb, J. W. (1991). Mechanism of rhodopsin kinase activation. J Biol Chem 266, 12949–12955. 42. Rim, J., Faurobert, E., Hurley, J. B., Oprian, D. D. (1997). In vitro assay for trans-phosphorylation of rhodopsin by rhodopsin kinase. Biochemistry 36, 7064–7070. 43. Yamamoto, S., Sippel, K., Berson, E. L., Dryja, T. P. (1997). Defects in the rhodopsin kinase gene in the Oguchi form of stationary night blindness. Nat Genet 15, 175–178.

136

Gross et al.

44. Yamamoto, S., Khani, S. C., Berson, E. L., Dryja, T. P. (1997). Evaluation of rhodopsin kinase gene in patients with retinitis pigmentosa. Exp Eye Res 65, 249–253. 45. Cideciyan, A. V., Zhao, X., Nielsen, L., Khani, S. C., Jacobson, S. G., Palczewski, K. (1998). Null mutation in the rhodopsin kinase gene slows recovery kinetics of rod and cone phototransduction in man. Proc Natl Acad Sci U S A 95, 328–333. 46. Dizhoor, A. M., Ray, S., Kumar, D., Niemi, G. A., Spencer, M., Brolley, D., Walsh, K. A., Philipov, P. P., Hurley, J. B. (1991). Recoverin: a calcium sensitive activator of retinal rod guanylate cyclase. Science 251, 915–918. 47. Makino, C. L., Dodd, R. L., Chen, J., Burns, M. E., Roca, A., Simon, M. I., Baylor, D. A. (2004). Recoverin regulates light-dependent phosphodiesterase activity in retinal rods. J Gen Physiol 123, 729–741. 48. Kawamura, S. (1993). Rhodopsin phosphorylation as a mechanism of cyclic GMP phosphodiesterase regulation by S-modulin. Nature 362, 855–857. 49. Gray-Keller, M. P., Polans, A. S., Palczewski, K., Detwiler, P. B. (1993). The effect of recoverin0like calcium-binding proteins on the photoresponse of retinal rods. Neuron 10, 523– 531. 50. Weiss, E. R., Ducceschi, M. H., Horner, T. J., Li, A., Craft, C. M., Osawa, S. (2001). Speciesspecific differences in expression of G-protein-coupled receptor kinase (GRK) 7 and GRK1 in mammalian cone photoreceptor cells: implications for cone cell phototransduction. J Neurosci 21, 9175–9184. 51. Tachibanaki, S., Arinobu, D., Shimauchi-Matsukawa, Y., Tsushima, S., Kawamura, S. (2005). Highly effective phosphorylation by G protein-coupled receptor kinase 7 of lightactivated visual pigment in cones. Proc Natl Acad Sci U S A 102, 9329–9334. 52. Liu, P., Osawa, S., Weiss, E. R. (2005). M opsin phosphorylation in intact mammalian retinas. J Neurochem 93, 135–144. 53. Sagi-Eisenberg, R., Traub, L. M., Spiegel, A. M., Zick, Y. (1989). Protein kinase C-mediated phosphorylation of retinal rod outer segment membrane proteins. Cell Signal 1, 519–531. 54. Inoue, M., Isayama, Y. (1984). Calcium ion and phospholipid-dependent protein kinase in rod outer segment. Jpn J Ophthalmol 28, 47–56. 55. Kapoor, C. L., Chader, G. J. (1984). Endogenous phosphorylation of retinal photoreceptor outer segment proteins by calcium phospholipid-dependent protein kinase. Biochem Biophys Res Commun 122, 1397–1403. 56. Kelleher, D. J., Johnson, G. L. (1985). Purification of protein kinase C from bovine rod outer segments. J Cyclic Nucleotide Protein Phosphor Res 10, 579–591. 57. Kelleher, D. J., Johnson, G. L. (1986). Phosphorylation of rhodopsin by protein kinase C in vitro. J Biol Chem 261, 4749–4757. 58. Kapoor, C. L., O’Brien, P. J., Chader, G. J. (1987). Phorbol ester- and light-induced endogenous phosphorylation of rat rod outer-segment proteins. Exp Eye Res 45, 545–556. 59. Wood, J. G., Hart, C. E., Mazzei, G. J., Girard, P. R., Kuo, J. F. (1988). Distribution of protein kinase C immunoreactivity in rat retina. Histochem J 20, 63–68. 60. Binder, B. M., Brewer, E., Bownds, M. D. (1989). Stimulation of protein phosphorylations in frog rod outer segments by protein kinase activators. Suppression of light-induced changes in membrane current and cGMP by protein kinase C activators. J Biol Chem 264, 8857–8864. 61. Newton, A. C., Williams, D. S. (1991). Involvement of protein kinase C in the phosphorylation of rhodopsin. J Biol Chem 266, 17725–17728. 62. Newton, A. C., Williams, D. S. (1993). Rhodopsin is the major in situ substrate of protein kinase C in rod outer segments of photoreceptors. J Biol Chem 268, 18181–18186.

Phosphorylation in Phototransduction

137

63. Williams, D. S., Liu, X., Schlamp, C. L., Ondek, B., Jaken, S., Newton, A. C. (1997). Characterization of protein kinase C in photoreceptor outer segments. J Neurochem 69, 1693–1702. 64. Greene, N. M., Williams, D. S., Newton, A. C. (1995). Kinetics and localization of the phosphorylation of rhodopsin by protein kinase C. J Biol Chem 270, 6710–6717. 65. Wolbring, G., Cook, N. J. (1991). Rapid purification and characterization of protein kinase C from bovine retinal rod outer segments. Eur J Biochem 201, 601–606. 66. Sokal, I., Hu, G., Liang, Y., Mao, M., Wensel, T. G., Palczewski, K. (2003). Identification of protein kinase C isozymes responsible for the phosphorylation of photoreceptor-specific RGS9-1 at Ser475. J Biol Chem 278, 8316–8325. 67. Zick, Y., Sagi-Eisenberg, R., Pines, M., Gierschik, P., Spiegel, A. M. (1986). Multisite phosphorylation of the alpha subunit of transducin by the insulin receptor kinase and protein kinase C. Proc Natl Acad Sci U S A 83, 9294–9297. 68. Adams, R. A., Liu, X., Williams, D. S., Newton, A. C. (2003). Differential spatial and temporal phosphorylation of the visual receptor, rhodopsin, at two primary phosphorylation sites in mice exposed to light. Biochem J 374, 537–543. 69. Phillips, W. J., Wong, S. C., Cerione, R. A. (1992). Rhodopsin/transducin interactions. II. Influence of the transducin-beta gamma subunit complex on the coupling of the transducinalpha subunit to rhodopsin. J Biol Chem 267, 17040–17046. 70. Udovichenko, I. P., Cunnick, J., Gonzales, K., Takemoto, D. J. (1993). Phosphorylation of bovine rod photoreceptor cyclic GMP phosphodiesterase. Biochem J 295(Pt 1), 49–55. 71. Udovichenko, I. P., Cunnick, J., Gonzalez, K., Takemoto, D. J. (1994). Functional effect of phosphorylation of the photoreceptor phosphodiesterase inhibitory subunit by protein kinase C. J Biol Chem 269, 9850–9856. 72. Wolbring, G., Schnetkamp, P. P. (1995). Activation by PKC of the Ca(2+)-sensitive guanylyl cyclase in bovine retinal rod outer segments measured with an optical assay. Biochemistry 34, 4689–4695. 73. Duda, T., Venkataraman, V., Krishnan, A., Sharma, R. K. (1998). Rod outer segment membrane guanylate cyclase type 1 (ROS-GC1) gene: structure, organization and regulation by phorbol ester, a protein kinase C activator. Mol Cell Biochem 189, 63–70. 74. Hu, G., Jang, G. F., Cowan, C. W., Wensel, T. G., Palczewski, K. (2001). Phosphorylation of RGS9-1 by an endogenous protein kinase in rod outer segments. J Biol Chem 276, 22287–22295. 75. Hu, G., Zhang, Z., Wensel, T. G. (2003). Activation of RGS9-1GTPase acceleration by its membrane anchor, R9AP. J Biol Chem 278, 14550–14554. 76. Baker, S. A., Martemyanov, K. A., Shavkunov, A. S., Arshavsky, V. Y. (2006). Kinetic mechanism of RGS9-1 potentiation by R9AP. Biochemistry 45, 10690–7. 77. Xiong, W., Nakatani, K., Ye, B., Yau, K. (1997). Protein kinase C activity and light sensitivity of single amphibian rods. J Gen Physiol 110, 441–452. 78. Jindrova, H., Detwiler, P. B. (1998). Protein kinase C and IP3 in photoresponses of functionally intact rod outer segments: constraints about their role. Physiol Res 47, 285–290. 79. Hamm, H. (1990). Regulation by light of cyclic nucleotide-dependent protein kinases and their substrates in frog rod outer segments. J Gen Physiol 95, 545–567. 80. Horner, T. J., Osawa, S., Schaller, M. D., Weiss, E. R. (2005). Phosphorylation of GRK1 and GRK7 by cAMP-dependent protein kinase attenuates their enzymatic activities. J Biol Chem 280, 28241–28250. 81. Wolbring, G., Schnetkamp, P. P. (1996). Modulation of the calcium sensitivity of bovine retinal rod outer segment guanylyl cyclase by sodium ions and protein kinase A. Biochemistry 35, 11013–11018. 82. Xu, L. X., Tanaka, Y., Bonderenko, V. A., Matsuura, I., Matsumoto, H., Yamazaki, A., Hayashi, F. (1998). Phosphorylation of the gamma subunit of the retinal photoreceptor cGMP

138

83. 84.

85.

86.

87. 88.

89.

90.

91.

92.

93. 94.

95.

96.

97. 98.

Gross et al. phosphodiesterase by the cAMP-dependent protein kinase and its effect on the gamma subunit interaction with other proteins. Biochemistry 37, 6205–6213. Ko, G. Y., Ko, M. L., Dryer, S. E. (2004). Circadian regulation of cGMP-gated channels of vertebrate cone photoreceptors: role of cAMP and Ras. J Neurosci 24, 1296–1304. Stella, S. L., Jr., Thoreson, W. B. (2000). Differential modulation of rod and cone calcium currents in tiger salamander retina by D2 dopamine receptors and cAMP. Eur J Neurosci 12, 3537–3548. Stella, S. L., Jr., Bryson, E. J., Thoreson, W. B. (2002). A2 adenosine receptors inhibit calcium influx through L-type calcium channels in rod photoreceptors of the salamander retina. J Neurophysiol 87, 351–360. Balasubramanian, N., Levay, K., Keren-Raifman, T., Faurobert, E., Slepak, V. Z. (2001). Phosphorylation of the regulator of G protein signaling RGS9–1 by protein kinase A is a potential mechanism of light- and Ca2+-mediated regulation of G protein function in photoreceptors. Biochemistry 40, 12619–12627. Lee, R. H., Brown, B. M., Lolley, R. N. (1990). Protein kinase A phosphorylates retinal phosducin on serine 73 in situ. J Biol Chem 265, 15860–15866. Bauer, P. H., Muller, S., Puzicha, M., Pippig, S., Obermaier, B., Helmreich, E. J., Lohse, M. J. (1992). Phosducin is a protein kinase A-regulated G-protein regulator. Nature 358, 73–76. Pagh-Roehl, K., Lin, D., Su, L., Burnside, B. (1995). Phosducin and PP33 are in vivo targets of PKA and type 1 or 2A phosphatases, regulators of cell elongation in teleost rod inner-outer segments. J Neurosci 15, 6475–6488. Thulin, C. D., Savage, J. R., McLaughlin, J. N., Truscott, S. M., Old, W. M., Ahn, N. G., Resing, K. A., Hamm, H. E., Bitensky, M. W., Willardson, B. M. (2001). Modulation of the G protein regulator phosducin by Ca2+/calmodulin-dependent protein kinase II phosphorylation and 14-3-3 protein binding. J Biol Chem 276, 23805–23815. Lee, R. H., Ting, T. D., Lieberman, B. S., Tobias, D. E., Lolley, R. N., Ho, Y. K. (1992). Regulation of retinal cGMP cascade by phosducin in bovine rod photoreceptor cells. Interaction of phosducin and transducin. J Biol Chem 267, 25104–25112. Yoshida, T., Willardson, B. M., Wilkins, J. F., Jensen, G. J., Thornton, B. D., Bitensky, M. W. (1994). The phosphorylation state of phosducin determines its ability to block transducin subunit interactions and inhibit transducin binding to activated rhodopsin. J Biol Chem 269, 24050–24057. Gaudet, R., Bohm, A., Sigler, P. B. (1996). Crystal structure at 2.4 angstroms resolution of the complex of transducin betagamma and its regulator, phosducin. Cell 87, 577–588. Gaudet, R., Savage, J. R., McLaughlin, J. N., Willardson, B. M., Sigler, P. B. (1999). A molecular mechanism for the phosphorylation-dependent regulation of heterotrimeric G proteins by phosducin. Mol Cell 3, 649–660. Kobayashi, Y., Hisatomi, O., Yamamoto, S., Tokunaga, F. (2002). Distribution of rodand cone-specific phosducins in retinas of non-mammalian vertebrates. Comp Biochem Physiol B Biochem Mol Biol 133, 77–83. Sokolov, M., Strissel, K. J., Leskov, I. B., Michaud, N. A., Govardovskii, V. I., Arshavsky, V. Y. (2004). Phosducin facilitates light-driven transducin translocation in rod photoreceptors. Evidence from the phosducin knockout mouse. J Biol Chem 279, 19149–19156. Chen, J., Yoshida, T., Nakano, K., Bitensky, M. W. (2005). Subcellular localization of phosducin in rod photoreceptors. Vis Neurosci 22, 19–25. Yamamoto, S., Hisatomi, O., Hasegawa, A., Kobayashi, Y., Tokunaga, F. (2007). Distribution and translocation of photoreceptor Gbetagamma-phosducin system in medaka retina. Photochem Photobiol 83, 35–39.

Phosphorylation in Phototransduction

139

99. Zhu, X., Craft, C. M. (2000). The carboxyl terminal domain of phosducin functions as a transcriptional activator. Biochem Biophys Res Commun 270, 504–509. 100. Zhu, X., Craft, C. M. (2000). Modulation of CRX transactivation activity by phosducin isoforms. Mol Cell Biol 20, 5216–5226. 101. Matsuura, I., Bondarenko, V. A., Maeda, T., Kachi, S., Yamazaki, M., Usukura, J., Hayashi, F., Yamazaki, A. (2000). Phosphorylation by cyclin-dependent protein kinase 5 of the regulatory subunit of retinal cGMP phosphodiesterase. I. Identification of the kinase and its role in the turnoff of phosphodiesterase in vitro [in process citation]. J Biol Chem 275, 32950–32957. 102. Hayashi, F., Matsuura, I., Kachi, S., Maeda, T., Yamamoto, M., Fujii, Y., Liu, H., Yamazaki, M., Usukura, J., Yamazaki, A. (2000). Phosphorylation by cyclin-dependent protein kinase 5 of the regulatory subunit of retinal cGMP phosphodiesterase. II. Its role in the turnoff of phosphodiesterase in vivo [in process citation]. J Biol Chem 275, 32958–32965. 103. Waldbillig, R. J., Fletcher, R. T., Chader, G. J., Rajagopalan, S., Rodrigues, M., LeRoith, D. (1987). Retinal insulin receptors. 2. Characterization and insulin-induced tyrosine kinase activity in bovine retinal rod outer segments. Exp Eye Res 45, 837–844. 104. Zick, Y., Spiegel, A. M., Sagi-Eisenberg, R. (1987). Insulin-like growth factor I receptors in retinal rod outer segments. J Biol Chem 262, 10259–10264. 105. Waldbillig, R. J., Fletcher, R. T., Somers, R. L., Chader, G. J. (1988). IGF-I receptors in the bovine neural retina: structure, kinase activity and comparison with retinal insulin receptors. Exp Eye Res 47, 587–607. 106. Bell, M. W., Desai, N., Guo, X. X., Ghalayini, A. J. (2000). Tyrosine phosphorylation of the alpha subunit of transducin and its association with src in photoreceptor rod outer segments [in process citation]. J Neurochem 75, 2006–2019. 107. Omri, B., Blancher, C., Neron, B., Marty, M. C., Rutin, J., Molina, T. J., Pessac, B., Crisanti, P. (1998). Retinal dysplasia in mice lacking p56lck. Oncogene 16, 2351–2356. 108. Molokanova, E., Krajewski, J. L., Satpaev, D., Luetje, C. W., Kramer, R. H. (2003). Subunit contributions to phosphorylation-dependent modulation of bovine rod cyclic nucleotide-gated channels. J Physiol 552, 345–356. 109. Savchenko, A., Kraft, T. W., Molokanova, E., Kramer, R. H. (2001). Growth factors regulate phototransduction in retinal rods by modulating cyclic nucleotide-gated channels through dephosphorylation of a specific tyrosine residue. Proc Natl Acad Sci U S A 98, 5880–5885. 110. Rajala, R. V., McClellan, M. E., Ash, J. D., Anderson, R. E. (2002). In vivo regulation of phosphoinositide 3-kinase in retina through light-induced tyrosine phosphorylation of the insulin receptor beta-subunit. J Biol Chem 277, 43319–43326. 111. Rajala, R. V., Anderson, R. E. (2001). Interaction of the insulin receptor beta-subunit with phosphatidylinositol 3-kinase in bovine ROS. Invest Ophthalmol Vis Sci 42, 3110–3117. 112. Fontaine, V., Kinkl, N., Sahel, J., Dreyfus, H., Hicks, D. (1998). Survival of purified rat photoreceptors in vitro is stimulated directly by fibroblast growth factor-2. J Neurosci 18, 9662–9672. 113. Valter, K., van Driel, D., Bisti, S., Stone, J. (2002). FGFR1 expression and FGFR1-FGF-2 colocalisation in rat retina: sites of FGF-2 action on rat photoreceptors. Growth Factors 20, 177–188. 114. Hallbook, F., Backstrom, A., Kullander, K., Ebendal, T., Carri, N. G. (1996). Expression of neurotrophins and trk receptors in the avian retina. J Comp Neurol 364, 664–676. 115. Okazawa, H., Kamei, M., Imafuku, I., Kanazawa, I. (1994). Gene regulation of trkB and trkC in the chick retina by light/darkness exposure. Oncogene 9, 1813–1818. 116. Di Polo, A., Cheng, L., Bray, G. M., Aguayo, A. J. (2000). Colocalization of TrkB and brain-derived neurotrophic factor proteins in green-red-sensitive cone outer segments. Invest Ophthalmol Vis Sci 41, 4014–4021.

140

Gross et al.

117. Nag, T. C., Wadhwa, S. (1999). Neurotrophin receptors (Trk A, Trk B, and Trk C) in the developing and adult human retina. Brain Res Dev Brain Res 117, 179–189. 118. Ko, G. Y., Ko, M. L., Dryer, S. E. (2001). Circadian regulation of cGMP-gated cationic channels of chick retinal cones. Erk MAP Kinase and Ca2+/calmodulin-dependent protein kinase II. Neuron 29, 255–66. 119. Hauck, S. M., Ekstrom, P. A., Ahuja-Jensen, P., Suppmann, S., Paquet-Durand, F., van Veen, T., Ueffing, M. (2006). Differential modification of phosducin protein in degenerating rd1 retina is associated with constitutively active Ca2+/calmodulin kinase II in rod outer segments. Mol Cell Proteomics 5, 324–336. 120. Fowles, C., Akhtar, M., Cohen, P. (1989). Interplay of phosphorylation and dephosphorylation in vision: protein phosphatases of bovine rod outer segments. Biochemistry 28, 9385–9391. 121. Selke, D., Anton, H., Klumpp, S. (1998). Serine/threonine protein phosphatases type 1, 2A and 2C in vertebrate retinae. Acta Anat (Basel) 162, 151–156. 122. Klumpp, S., Selke, D. (2000). Purification and characterization of protein phosphatase type 2C in photoreceptors. Methods Enzymol 315, 570–578. 123. Klumpp, S., Selke, D., Fischer, D., Baumann, A., Muller, F., Thanos, S. (1998). Protein phosphatase type-2C isozymes present in vertebrate retinae: purification, characterization, and localization in photoreceptors. J Neurosci Res 51, 328–338. 124. Palczewski, K., Hargrave, P. A., McDowell, J. H., Ingebritsen, T. S. (1989). The catalytic subunit of phosphatase 2A dephosphorylates phosphoopsin. Biochemistry 28, 415–419. 125. Palczewski, K., McDowell, J. H., Jakes, S., Ingebritsen, T. S., Hargrave, P. A. (1989). Regulation of rhodopsin dephosphorylation by arrestin. J Biol Chem 264, 15770–15773. 126. King, A. J., Andjelkovic, N., Hemmings, B. A., Akhtar, M. (1994). The phospho-opsin phosphatase from bovine rod outer segments. An insight into the mechanism of stimulation of type-2A protein phosphatase activity by protamine. Eur J Biochem 225, 383–394. 127. Sokal, I., Hu, G., Liang, Y., Mao, M., Wensel, T. G., Palczewski, K. (2003). Identification of protein kinase C isozymes responsible for the phosphorylation of photoreceptor-specific RGS9-1 at Ser475. J Biol Chem 278, 8316–25. 128. Brown, B. M., Carlson, B. L., Zhu, X., Lolley, R. N., Craft, C. M. (2002). Light-driven translocation of the protein phosphatase 2A complex regulates light/dark dephosphorylation of phosducin and rhodopsin. Biochemistry 41, 13526–13538. 129. Lyons, B. L., Smith, R. S., Hurd, R. E., Hawes, N. L., Burzenski, L. M., Nusinowitz, S., Hasham, M. G., Chang, B., Shultz, L. D. (2006). Deficiency of SHP-1 protein-tyrosine phosphatase in “viable motheaten” mice results in retinal degeneration. Invest Ophthalmol Vis Sci 47, 1201–1209. 130. Bell, M. W., Alvarez, K., Ghalayini, A. J. (1999). Association of the tyrosine phosphatase SHP-2 with transducin-alpha and a 97-kDa tyrosine-phosphorylated protein in photoreceptor rod outer segments. J Neurochem 73, 2331–2340. 131. Okada, T., Sugihara, M., Bondar, A. N., Elstner, M., Entel, P., Buss, V. (2004). The retinal conformation and its environment in rhodopsin in light of a new 2.2 A crystal structure. J Mol Biol 342, 571–583. 132. Kisselev, O. G., McDowell, J. H., Hargrave, P. A. (2004). The arrestin-bound conformation and dynamics of the phosphorylated carboxy-terminal region of rhodopsin. FEBS Lett 564, 307–311. 133. Ames, J. B., Levay, K., Wingard, J. N., Lusin, J. D., Slepak, V. Z. (2006). Structural basis for calcium-induced inhibition of rhodopsin kinase by recoverin. J Biol Chem 281, 37237–37245.

7 The cGMP Signaling Pathway in Retinal Photoreceptors and the Central Role of Photoreceptor Phosphodiesterase (PDE6) Rick H. Cote CONTENTS Overview of Cyclic Guanosine Monophosphate Signaling Pathways The Cellular Context of cGMP Signaling in Vertebrate Retinal Photoreceptors Photoreceptor PDE (PDE6) Structure and Function PDE6 Regulation Conclusions References

OVERVIEW OF CYCLIC GUANOSINE MONOPHOSPHATE SIGNALING PATHWAYS All eukaryotic cells utilize cyclic nucleotides—specifically cyclic adenosine monophosphate (cAMP) and cyclic guanosine monophosphate (cGMP)—as intracellular messengers in a wide variety of cell signaling pathways. In addition to the visual signaling pathway (the focus of this review), cGMP is involved in numerous other physiological processes, including vascular smooth muscle relaxation, natriuresis, platelet function, neutrophil adhesion, sperm motility, neuronal signaling, and other sensory transduction systems [1]. The metabolism of cGMP is controlled by the synthetic enzymes, guanylate cyclases (GCs), and hydrolytic enzymes, cyclic nucleotide phosphodiesterases (PDEs). Cytoplasmic levels of cGMP may also be modulated nonenzymatically by sequestration by cGMP-binding proteins [2] or by transport mechanisms that cause cGMP efflux from the cell [3]. Changes in cytoplasmic cGMP concentration affect cGMP signaling pathways by changing the extent of binding to specific cGMP-binding proteins (receptors). Targets of cGMP action include cGMPdependent protein kinases (PKGs), cyclic nucleotide-gated (CNG) ion channels, and cGMP-binding PDEs, all of which are allosterically regulated by cGMP binding to noncatalytic regulatory sites on these proteins.

From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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Regulation of Intracellular cGMP Levels in Photoreceptor Cells Guanylate cyclase (GC) catalyzes the synthesis of cGMP from guanosine triphosphate (GTP). Vertebrates have two major families of GC, soluble and membrane associated. The two membrane-associated GCs found in photoreceptor cells (GC-2E and GC-2 F; also abbreviated as ROS-GCs or Ret-GCs in the literature) consist of an extracellular domain (of unknown function), a single-pass transmembrane segment, a kinase-homology domain, and a catalytic domain [4, 5]. The photoreceptor GCs are not regulated by binding of ligands to the extracellular domain as is the case for several other membrane-associated GCs [5]. Instead, photoreceptor GCs are regulated in a calcium-dependent manner by three distinct GC-activating proteins (GCAPs; GCAP1, GCAP2, and GCAP3 [4, 6, 7]). The breakdown of cyclic nucleotides in cells is catalyzed by cyclic nucleotide PDEs. In vertebrates, there are 11 families of PDEs that share a conserved catalytic domain but differ in their substrate specificity (cAMP-, cGMP-, or dual-specific), regulatory mechanisms, and pharmacological sensitivity [8]. Rods and cones express a photoreceptor-specific PDE named PDE6 that has a very high catalytic efficiency when activated by light, is regulated by its inhibitory γ-subunit, and shares structural and pharmacological sensitivity with PDE5, a PDE abundant in vascular smooth muscle and other tissues [9]. In addition to metabolic regulation of cGMP by GC and PDE activities, the free cytoplasmic cGMP concentration can also be regulated by cGMP transport out of the cell or by cGMP sequestration (i.e., binding to specific binding sites) within the cell. Transport systems have been identified that selectively pump cGMP out of various cell types [3, 10, 11], but to date evidence for cGMP efflux from photoreceptors is lacking. In contrast, high-affinity cGMP-binding proteins (e.g., PDE6 itself) are present in photoreceptor cells and are likely to contribute substantially to reducing the free cGMP concentration [2]. Sequestration of cGMP is indeed a major factor in determining the cytoplasmic free cGMP concentration of 2–4 µM (inferred from electrophysiological studies; [12, 13]) since the total cGMP concentration in the rod outer segment is tenfold higher [14]. Downstream Targets of cGMP Action in Photoreceptor Cells All of the above-mentioned downstream targets of cGMP are present in photoreceptor cells: the PKG, the CNG ion channels, and the cGMP-binding PDE6. cGMP-Dependent Protein Kinase There is not much known about the abundance of PKG in photoreceptor cells, potential protein substrates for reversible phosphorylation, or the relevance of PKG for the phototransduction pathway. Some evidence indicates that cAMP-dependent protein kinase (PKA) is more prevalent than PKG [15, 16], and other studies do not unequivocally distinguish PKG from PKA [17, 18]. Considering the relatively high cGMP levels in photoreceptor cells, the potential for cGMP to bind to and activate PKA (i.e., “cross-talk”) cannot be ruled out. The observation that several PKG/PKA substrates undergo light-dependent phosphorylation/ dephosphorylation supports the idea that cyclic nucleotide-dependent kinases may be involved in some aspects of visual signaling. For example, light-dependent dephosphorylation of components I and II (in amphibian photoreceptors) and phosducin (in mammalian and fish photoreceptors) is consistent with the idea that the light-induced drop in free cGMP levels results in PKA/PKG inactivation and thereby causes dephosphorylation of

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the above-mentioned phosphoproteins [19–25]. The abilities of these dephosphorylated proteins to preferentially bind to transducin βγ dimers [25–27] and to assist in lightdependent protein translocation within the photoreceptor cell [28] are consistent with a role for PKG/PKA in long-term light adaptation. Cyclic Nucleotide-Gated Ion Channels The CNG ion channels in rod and cone cells belong to a large superfamily of ion channels that share a similar structure, including six transmembrane segments that selfassociate to constitute the pore of the ion channel [29]. The rod and cone CNG channels are heterotetramers (rod, 3 CNGA1 and 1 CNGB1; cone, 2 CNGA3, 2 CNGB3 [30, 31]). Activation (opening) of the CNG ion channel results from a highly cooperative binding of four cyclic nucleotide molecules to the C-terminal cyclic nucleotide-binding domain in the channel [32]. Channel closure in the plasma membrane resulting from the drop in cGMP levels induces hyperpolarization of the cell membrane and the generation of the receptor potential. Rod and cone CNG channels are optimized to instantaneously sense and respond to changes in cGMP concentration induced by activation of the visual excitation pathway. Rapid responsiveness to fluctuations in cGMP levels reflects the relatively low affinity of cGMP binding along with fast dissociation and association rates for the cGMP-binding sites. The cooperativity (Hill coefficient ~3) with which four cGMP molecules bind amplifies small changes in cGMP concentration [33]. An important aspect of the ionic permeability of the CNG channel is its relatively high calcium permeability; this generates a calcium feedback signal (in concert with a Na+/K+-Ca2+ exchanger) when the channels close during visual excitation, thus allowing for calcium-dependent reactions involved in recovery and adaptation to occur [29, 33]. Calcium regulatory proteins (calmodulin in rods, an uncharacterized calcium-binding protein in cones) bind to and regulate the photoreceptor CNG channel by reducing its cGMP sensitivity. Desensitization of CNG channels by reducing cGMP-binding affinity is also reported to be affected by tyrosine or serine/threonine phosphorylation, diacylglycerol or related metabolites, and retinoids [29, 32, 34–36], but the physiological significance of these regulatory mechanisms is uncertain. PDE6 Is a High-Affinity cGMP-Binding Protein In addition to having a catalytic domain responsible for lowering cGMP levels during visual excitation, PDE6 contains a regulatory domain consisting of two tandemly arrayed domains that bind cGMP with high affinity. These cGMP-binding sites serve to sequester a majority of the total cGMP in the photoreceptor cell as well as having regulatory properties, both of which are discussed in detail in this chapter. THE CELLULAR CONTEXT OF cGMP SIGNALING IN VERTEBRATE RETINAL PHOTORECEPTORS Compartmentation of cGMP Signaling in Photoreceptor Outer Segments Vertebrate rod and cone photoreceptor cells are specialized neurons consisting of several functionally and structurally distinct cellular compartments: (1) The phototransducing outer segment portion of the cell contains densely packed membranes optimized for

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photon capture, signal transduction, and the initial membrane hyperpolarization. (2) The nonmotile “connecting cilium” region connects the outer segment to the inner segment and actively regulates the transport of proteins and metabolites between these two compartments. (3) The inner segment is comprised of the metabolic machinery and is itself compartmentalized: Mitochondria are concentrated in the “ellipsoid” region nearest the connecting cilium, whereas organelles dedicated to protein biosynthesis are located between the mitochondria-rich ellipsoid region and the cell nucleus. (4) The synaptic terminals of rods and cones tonically release neurotransmitters in the dark and respond to membrane hyperpolarization by suppressing synaptic vesicle release in the light. The photoreceptor synapse communicates with second-order bipolar and horizontal cells to propagate the photoresponse. (For reviews, see [37, 38].) Physiology of the Photoreceptor Response to Light In the dark, a circulating “dark current” is maintained by entry of sodium and calcium through open CNG ion channels in the outer segment plasma membrane; this is concurrent with the extrusion of sodium by a Na+/K+-ATPase (adenosine triphosphatase) and the efflux of potassium by K+ channels (both localized to the inner segment). A Na+/Ca2+-K+ exchanger (in the outer segment) and other ion channels in the inner segment further regulate ion conduction and transport in photoreceptors. On illumination, this dark current is interrupted, and the change in membrane potential that results from channel closure is passively propagated from the outer segment through the inner segment to the synaptic terminal [39–41]. Remarkably, rod photoreceptors can detect individual photons and can generate discrete, reproducible photoresponses from these single-photon events [42]. In contrast, cone photoreceptors are less light sensitive than rods; their photoresponses are smaller and faster than for rods, but they operate over an enormous range of ambient light intensities. The rising phase of the photoresponse (termed visual excitation) is dominated by the kinetics of activation of the cGMP signaling pathway (discussed in detail in the next section). However, to rapidly respond to changes in light stimuli, the visual excitation process must be rapidly terminated. Photoresponse recovery is tightly coordinated with the components of the cGMP excitation pathway but also depends on another second messenger, calcium, to control the kinetics of the recovery to the dark-adapted state. Furthermore, photoreceptors also undergo “light adaptation” in the presence of background illumination, a process that allows rods and cones to increase the range of light intensities over which visual transduction can operate. On exposure to background illumination, the sensitivity of rods and cones to flash stimulation is decreased, resulting in smaller photoresponses that have faster recovery kinetics. Calcium plays a central role in the underlying mechanisms of light adaptation, acting through calcium regulatory proteins to modulate several steps in the cGMP signaling pathway. (For reviews, see [40, 43–47].) Biochemical Cascade of Visual Excitation The phototransduction cascade is a prototypical heterotrimeric G protein-coupled signaling pathway (Fig. 1). The excitation process is triggered by absorption of a photon by the visual receptor rhodopsin, which activates a photoreceptor-specific G protein, transducin, that then activates the effector enzyme PDE6. The resultant drop in

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Fig. 1. The cyclic guanosine monophosphate (cGMP) signaling pathway for visual excitation in vertebrate rod photoreceptors. Top: In dark-adapted rod photoreceptors, cytoplasmic cGMP (small gray circle) and calcium concentrations are high, and some of the cGMP-gated cation channels in the plasma membrane are fully liganded with cGMP and in their open state. This permits entry of Na+ and Ca2+ through the pore. Rhodopsin, transducin, and phosphodiesterase 6 (PDE6) are in their nonactivated states. Bottom: On absorption of a photon by rhodopsin, isomerization of the 11-cis retinal chromophore causes receptor activation (R*). This leads to binding of transducin to R*, guanine nucleotide exchange of guanosine diphosphate (GDP) for guanosine triphosphate (GTP), and formation of the activated transducin α-subunit with bound GTP (Tα*). The Tα* species then binds PDE6 holoenzyme, causing deinhibition by the γ-subunit (Tα*-P*) and a large acceleration of catalysis of cGMP to 5’-GMP at the active site. The light-induced drop in cGMP concentration induces the ligand-gated ion channel to close, causing membrane hyperpolarization. Ongoing extrusion of calcium by the Na+/Ca2+-K+ exchanger in the absence of calcium influx through the channel also causes intracellular calcium concentration to decline (which is vital for the recovery process).

second-messenger concentration (i.e., cGMP) leads to dissociation of cGMP bound to the CNG ion channel, closure of the ion channel, and membrane hyperpolarization. Central Components of the cGMP Signaling Pathway The first step in vertebrate vision is the photoisomerization of the retinal chromophore (11-cis retinal) of the visual pigment on the outer segment membrane (Fig. 1).

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Rhodopsin is a member of the G protein-coupled receptor superfamily, in which 11-cis retinal locks rhodopsin into its inactive conformation in the dark. Photoactivation causes isomerization of 11-cis retinal to all-trans retinal. This causes movement of the transmembrane α-helices that surround the chromophore, producing metarhodopsin II, the activated form of the receptor [48, 49]. Conformational changes in the C-terminal tail and cytoplasmic loops of metarhodopsin II allow rhodopsin to bind with high affinity to the heterotrimeric G protein, transducin [50]. This interaction catalyzes the exchange of bound GDP for GTP on the transducin α-subunit, causing dissociation of the α-subunit (with bound GTP) from the βγ dimer [51]. Activated transducin then binds to its effector in this signaling cascade, PDE6, displacing the inhibitory PDE6 γ-subunit and accelerating the catalysis of cGMP (see the section “Transducin Activation of Rod PDE6 During Visual Excitation” for details). Because of the significant lifetime of metarhodopsin II, 20–100 transducin molecules (and hence PDE6 molecules) can be activated per photoisomerization event [52, 53]; this represents the first stage of amplification of the signaling cascade. A second stage in signal amplification follows stoichiometric activation of PDE6 by transducin: Each activated PDE6 can break down many thousands of cGMP molecules per second (see the section “PDE6 Has Evolved to Meet the Special Demands of the Central Effector of Visual Transduction”). The overall gain of this amplified excitation pathway is well over 100,000 cGMP molecules hydrolyzed per activated rhodopsin [51]. The very rapid drop in cytoplasmic cGMP levels on light activation ensures that cGMP dissociates quickly from binding sites on the CNG ion channel in the plasma membrane. This sequence leading from photoisomerization of visual pigment to channel closure and membrane hyperpolarization constitutes the set of reactions defined as visual excitation. Termination and Adaptation of the Light Response The recovery of the dark-adapted state following illumination occurs in a precisely controlled manner that optimizes both the temporal resolution of visual stimuli as well as the ability of photoreceptors to light adapt over the 1012 range of photic stimuli on earth. Each step in the visual excitation pathway must be deactivated to restore cGMP levels and to return the components of the excitation pathway to their inactive states. The recovery of the photoresponse depends on inactivation of the metarhodopsin II state of rhodopsin as well as deactivation of GTP-bound form of the transducin α-subunit. Because the kinetics of the recovery phase of the light response are highly stereotypical, it has been appreciated for some time that a single deactivation step must be rate limiting [43]. Using a transgenic approach, it has been shown that the rate of GTP hydrolysis by transducin α-subunit represents the rate-limiting step of rod photoresponse recovery [53]. By regulating the guanosine triphosphatase (GTPase) rate of activated transducin, the lifetime of activated PDE6 is thereby precisely controlled (see section “Deactivation of Transducin”). Light adaptation extends the operating range of the photoresponse of rods and cones by dampening the response amplitude in response to an incremental change in light intensity as well as accelerating the kinetics of the photoresponse. Whereas visual excitation requires consideration only of one second messenger (i.e., cGMP), calcium plays a central role in many aspects of photoreceptor adaptation. The cytoplasmic concentration of calcium

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in dark-adapted photoreceptor outer segments (400–600 nM) rapidly decreases on light exposure to 10–50 nM as a consequence of channel closure concomitant with continued extrusion of calcium by the Na+/Ca2+-K+ exchanger. Calcium regulates light adaptation primarily through three mechanisms: regulation of GC activity, regulation of the rate of inactivation of rhodopsin by rhodopsin kinasemediated phosphorylation, and modulation of the affinity cGMP-gated ion channel for cGMP. Each process is regulated by distinct calcium-binding proteins: GC-activating proteins (GCAPs) for GC, recoverin/S-modulin for rhodopsin kinase, and calmodulin for the ion channel. Of these three calcium-dependent control steps, the dominant one for light adaptation is the regulation of GC activity [44–46, 54]. Deactivation of Rhodopsin Metarhodopsin II inactivation involves phosphorylation by a specific G protein-coupled receptor kinase (GRK1 in rods, GRK7 in human cones) with activity that is regulated by the calcium-binding protein recoverin/S-modulin [55–57]. Calcium-recoverin binds to rhodopsin kinase and inhibits its ability to phosphorylate activated rhodopsin, thereby prolonging the activated state of the receptor [58]. Once phosphorylated, arrestin binds to phosphorylated rhodopsin to complete the inactivation process [59]. Pigment regeneration of the photobleached chromophore requires enzymatic and transport reactions, termed the retinoid cycle [60]. Deactivation of Transducin The inactivation of the α-subunit of transducin requires hydrolysis of bound GTP. The intrinsic GTPase rate of transducin is slow but can be accelerated when complexed with the regulator of G protein signaling 9 (RGS9 [61]), the type 5 G-protein β-subunit [62], and the RGS9 anchor protein (R9AP [63]). The PDE6 γ-subunit plays an elegant negative-feedback role by binding to RGS9 and enhancing the affinity of the RGS9 protein complex for α*-GTP (α-subunit with GTP bound) [64–66], thereby accelerating its intrinsically slow GTPase activity. This role of the γ-subunit to bind RGS9 serves to turn off PDE6 activation in a precisely timed manner that is critical to the kinetics of the recovery process. Importantly, this GTPase accelerating role of the γ-subunit does not interfere with the ability of α*-GTP to efficiently activate PDE6 during the initial stage of visual excitation [51]. Deactivation of PDE6 PDE6 inactivation occurs when the PDE6 γ-subunit is released from its binding site on the deactivated α-subunit of transducin and reinhibits the enzyme’s active site. The strength of the interaction of the γ-subunit for transducin versus PDE6 is modulated by the state of occupancy of cGMP at the PDE6 regulatory GAF domains. This feedback regulation mechanism is discussed in detail in the section “Functions of the Regulatory cGMP-Binding GAF Domains of PDE6”. A nonenzymatic mechanism for restoring cGMP levels by utilizing cGMP bound to high-affinity sites on the PDE6 GAF domains [67] is theoretically possible [2] but has not been experimentally supported [68], in large part because the catalytic power of activated PDE6 will hydrolyze cGMP as it dissociates from its binding sites.

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Other mechanisms for inactivating PDE6 independent of the hydrolysis of GTP by transducin α-subunit have been proposed [69–71]. For example, free γ-subunit (i.e., not associated with transducin or PDE6) has been shown to reinhibit transducin-activated PDE6 in vitro even though the transducin α-subunit remains persistently activated [72, 73]. Using a transgenic animal overexpressing the γ-subunit further supports the idea that the free γ-subunit is able to bind to and inhibit transducin-activated PDE6 [71], but the physiological significance of this is unclear since the concentration of γ-subunit in rod outer segments is equal to the concentration of PDE6 catalytic subunits, and all γ-subunit is found membrane associated [73]. Other potential regulatory mechanisms for PDE6 involving novel binding proteins are considered in the section Potential PDE6 Regulatory Binding Proteins. Activation of GC Rapid restoration of cGMP levels following PDE6 deactivation requires calciumdependent activation of photoreceptor GCs by their GCAPs. When the cytoplasmic concentration drops below about 500 nM, calcium dissociates from the GCAPs, relieving inhibition of GC and accelerating cGMP synthesis [4, 6, 7]. The powerful calcium feedback mechanism involving GCAP-GC regulation of cGMP synthesis helps determine the amplitude and temporal characteristics of the photoresponse and regulates the cGMP metabolic flux that is modulated during light adaptation [54]. Regulation of the CNG Ion Channel In rod photoreceptors, calcium/calmodulin binding to the β-subunit of the CNG channel serves to decrease the cGMP sensitivity of the channel [74]. In cone photoreceptors, modulation of cGMP sensitivity by calcium is greater than in rods, but the identity of calcium regulatory protein is not certain [75, 76]. PHOTORECEPTOR PDE (PDE6) STRUCTURE AND FUNCTION The Cyclic Nucleotide Phosphodiesterase Superfamily The photoreceptor PDE6 enzyme found in retinal rods and cones is a member of the class I PDE superfamily, of which 11 distinct gene families exist in vertebrates (Fig. 2). Class I PDEs all contain the characteristic PDEase I catalytic domain. The 11 PDE families are readily distinguished by comparison of their primary amino acid sequences as well as by their regulatory mechanisms, substrate preference, pharmacological inhibitor specificity, and expression patterns [8]. Three PDE families (PDE5, PDE6, and PDE9) strongly prefer cGMP as the substrate, three are cAMP specific (PDE4, PDE7, PDE8), and the rest do not discriminate between the two substrates. Five PDE families (PDE2, PDE5, PDE6, PDE10, PDE11) contain two tandem GAF domains in their N-terminal regulatory domain, which, in the case of PDE6, bind cGMP with high affinity [77]. Other PDE families are subject to regulation by calcium/calmodulin, phosphorylation, and extrinsic regulatory proteins [8]. The photoreceptor PDE6 is most closely related to PDE5 in its amino acid sequence, substrate preference, and pharmacological profile but differs in its mechanism of regulation [9].

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Fig. 2. The superfamily of cyclic nucleotide phosphodiesterases (PDEs). In humans, there are 21 PDE genes that are grouped into 11 different families, PDE1 though PDE11. Many of the PDE families exist as multiple splice variants (not shown), although this is not the case for PDE6. All class I PDEs share a highly homologous catalytic domain (PDEase I) in the C-terminal region of the linear sequence. The N-terminal half contains regulatory elements and targeting elements (defined in the box). The PDE6 family is unique in having a C-terminal membrane-targeting domain and in being directly regulated by an inhibitory γ-subunit (not shown). Abbreviations: PAS, Per-Arnt-Sim; GAF, see text; Cam, Calmodulin

Subunit Composition of Rod and Cone PDE6 Holoenzyme Although all class I PDEs are believed to exist as a dimer of two catalytic subunits, rod PDE6 is the only one in which the catalytic dimer consists of two nonidentical α (PDE6A) and β (PDE6B) subunits [78–81]. In contrast, cone PDE6 is a homodimeric enzyme composed of α′ (PDE6C) subunits [82, 83]. A second distinctive trait of PDE6 (this one shared by rod and cone isoforms) is the association of high-affinity inhibitory rod or cone γ-subunits to the corresponding catalytic dimer [84, 85]. The rod PDE6 holoenzyme (Fig. 3A) has been conclusively shown to be composed of two γ-subunits bound to the rod αβ-dimer [80, 86], and the cone enzyme is also assumed to be a heterotetramer. Catalytic Subunit The primary sequence of the α, β, and α′ catalytic subunits of PDE6 [87–89] consist of an N-terminal region of unknown function, two regulatory GAF domains (GAFa and GAFb) arranged in tandem, the catalytic domain, and a C-terminal motif that is subject to isoprenylation (Fig. 3B). Regulatory GAF Domain The tandem GAF domains (named for their occurrence in cGMP binding PDEs, certain adenylate cyclases, and the Escherichia coli FhlA protein [77, 90]) serve several

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Fig. 3. Domain organization of the catalytic and inhibitory subunits of phosphodiesterase 6 (PDE6). A The rod PDE6 catalytic dimer (left) consists of nonidentical α- and β-subunits that are farnesylated (α-subunit) or geranylgeranylated (β-subunit). The enzyme-active site is denoted by the notch, while the cGMP-binding site is indicated by a circular pocket. Right: The γ-subunit binds to both the active site (to inhibit catalysis) and to the GAF domains (to stabilize cyclic guanosine monophosphate [cGMP] binding). B The ~100-kDa catalytic subunit of PDE6 consists of two tandem GAF domains, a catalytic domain, and a prenylated C-terminus. C The 10-kDa γ-subunit has several functional domains: a proline-rich (Pro-rich) region and a polycationic region (PC region) that both interact with the cGMP-binding site on PDE6, an α-helical region, and the C-terminal residues (CT), which bind to the active site of PDE6. Activated transducin α-subunit also interacts with the γ-subunit at both its PC region and the α-helical region.

important functions. First, the GAF domains of PDE6 contain nucleotide-binding pockets (distinct from the enzyme’s active sites) that bind cGMP with high affinity [91]. The PDE6 GAF domains also serve a function unique to the PDE6 family, namely, to bind the inhibitory γ-subunit in a cGMP-dependent manner. Finally, the GAF domains of PDE6 are responsible for catalytic subunit dimerization [92, 93]. Although the rod PDE6 holoenzyme (αβγ2) has two GAF domains per catalytic subunit, rod PDE6 binds only two cGMP molecules with high affinity [94–96]. A second class of low-affinity cGMP-binding sites found in rod outer segments [97] might also represent cGMP binding to the remaining PDE6 GAF domains of the catalytic dimer, but this has not been experimentally verified to date. The high-affinity cGMP-binding sites on PDE6 catalytic subunits discriminate cGMP over cAMP by about 106-fold [98, 99]. The cGMP-binding site in PDE6 has been localized to the GAFa domain in rod and cone

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isoforms, and several amino acid residues important for high-affinity interactions with cGMP have been defined [99, 100]. The GAFa domain is also a major binding site for the inhibitory γ-subunit of PDE6, as judged by cross-linking and site-directed mutagenesis work [100–102]. Differences in the binding of the γ-subunit to the α- and β-subunit reveal potential structural differences between α-GAFa and β-GAFa that may account for heterogeneity in cGMP or γ-subunit affinity for the holoenzyme [102]. The proximity of γ-subunit interacting sites with the cGMP-binding pocket in GAFa provides structural support for the positive cooperativity between cGMP and γ-subunit binding (discussed in section “Functions of the Regulatory cGMP-binding GAF domains of PDE6”). The GAFa domains of rod and cone PDE6 catalytic subunits serve as the primary dimerization domain between two individual subunits. This is evident in the structural model of the PDE6 holoenzyme [103]. The N-terminal region of the GAFa was shown by a mutagenesis study to be essential for dimerization of rod PDE6 [92]; these studies also support the view that rod PDE6 exists primarily or exclusively as αβ heterodimers, consistent with earlier biochemical evidence [81]. Catalytic Domain The PDE6 catalytic domain contains the same invariant catalytic site residues that typify all class I PDEs, referred to as PDEase_I in the Conserved Domain Database (CDD) at the National Center for Biotechnology Information (NCBI). Multiple-sequence alignment of the catalytic domains of three human rod and cone PDE6 genes and the PDE5 gene reveals 84% amino acid identity between the PDE6A and PDE6B rod subunit isoforms, about 75% amino acid identity between rod and cone PDE6C subunits, and 43% identity between PDE5 and PDE6. Because of the difficulty in expressing functional PDE6 catalytic subunits in heterologous systems [104–108], site-directed mutagenesis of PDE6 has not been feasible. Instead, progress in understanding the structure and function of the PDE6 catalytic domain has relied on constructing chimeric proteins containing both PDE5 and PDE6 sequences in conjunction with structural homology modeling with known catalytic domain crystal structures [105]. This approach has identified some of the important residues responsible for the very high turnover number (kcat) of PDE6 compared to PDE5 [108, 109]. The histidine residues responsible for binding divalent cations in the active site of PDE5 [110, 111] are present in PDE6 and bind zinc with high affinity and magnesium with lower affinity [112]. These divalent cations are not only critical for the catalytic mechanism but also confer structural stability to the enzyme. As mentioned, the nonactivated PDE6 enzyme is inhibited by direct binding of its inhibitory γ-subunit. The γ-subunit exerts its inhibitory action by directly binding to the catalytic pocket, thereby blocking access of cGMP to its binding site [113, 114]. PDE5/6 chimeras have also proved useful in identifying specific γ-interacting amino acid residues (PDE6 α′: M758, Q752, F777, and F781) within the M loop of the catalytic domain structure [108, 109, 115]. C-Terminal Prenylation Although several PDE families contain signaling motifs that target the enzyme to the membrane, PDE6 is the only vertebrate PDE family that is either farnesylated (PDE6A) or geranylgeranylated (PDE6B and, probably, PDE6C) at a C-terminal CAAX (C = cysteine,

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A = aliphatic, X = any amino acid) motif [116, 117]. The prenylated, carboxymethylated C-termini are responsible for anchoring PDE6 to the outer segment disk membrane [118], thereby facilitating two-dimensional collisions with transducin during visual excitation. PDE6 is membrane bound, except when the 17-kDa PrBP/δ protein (originally termed PDEδ) is present to bind to the prenyl groups and solubilize PDE6 from the membrane (see section “Potential PDE6 regulatory binding proteins” for discussion). Disk membrane association of PDE6 can be disrupted in vitro by altering the ionic, nucleotide, divalent cation, or illumination conditions, which is useful for purification of the enzyme [119]. Inhibitory γ-Subunit The γ-subunit of rod and cone PDE6 [120, 121] serves a remarkable number of functions considering its small size (~10 kDa). Most likely, the γ-subunit exists in solution as an unfolded or intrinsically disordered protein due to its minimal secondary structure [122, 123]. This may account for the ability of this small protein to span the distance from the GAFa domain to the active site on the catalytic domain of the catalytic dimer [102]. Important regions of the γ-subunit include the following: 1. A proline-rich region (amino acids [a.a.] 22–28 of the rod sequence) is a potential site of interaction with proteins containing SH3 (src homology3) domains [124] as well as a site for protein phosphorylation by proline-directed kinases [125–127]. 2. The polycationic region (a.a. 29–45) is a major site of interaction with the transducin α-subunit during light activation of PDE6 [128–133], as well as a substrate for phosphorylation by serine/threonine kinases [127, 134, 135] and adenosine diphosphate (ADP) ribosylation at two arginine residues [136, 137]. 3. The combined proline-rich and polycationic regions of the γ-subunit is the strongest site of interaction with the PDE6 catalytic dimer [102, 128, 138–142] and enhances the binding of cGMP to the regulatory GAF domains [142, 143]. 4. The region between the polycationic and α-helical segments of the γ-subunit contains a site of interaction (a.a. 66) with RGS9-1 [66], which serves to accelerate the GTPase activity of activated transducin [61, 64, 66]. 5. An α-helical region of the γ-subunit (a.a. 62–83) is a major site of interaction with activated transducin α-subunit and is primarily responsible for the GTPase accelerating activity of the γ-subunit [66, 144–150]. In addition, amino acids F73, N74, H75, and L78 of the γ-subunit interact with the PDE6 catalytic subunits [102, 150], even though the α-helical region itself does not directly block the active site. It is thus likely that transducin binds to one face of the γ-subunit α-helical region and disrupts contacts between the opposite face of the α-helical domain and the catalytic domain of PDE6. 6. The last five C-terminal residues (a.a. 83–87) directly bind to the active site to cover the catalytic pocket and block cyclic nucleotide entry [113–115, 128, 131, 146]. It must also be emphasized that the γ-subunit interacts differently with the α- and β-subunits of rod PDE6, as judged by both biochemical and structural evidence [73, 96, 102, 142, 151]. Because the GAFa domains of the α- and β-subunits of rod PDE6 are more dissimilar than the GAFb or catalytic domains, it is possible that differences in γ-subunit binding affinity or functional properties may result from differences in interactions of γ with the GAFa domains.

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PDE6 Has Evolved to Meet the Special Demands of the Central Effector of Visual Transduction The exquisite light sensitivity and temporal resolution of the visual signaling pathway in retinal photoreceptors requires precise regulation of PDE6 in rod and cone outer segments. On the one hand, spontaneous activation of PDE6 must be minimized in the dark-adapted state to enhance the absolute light sensitivity and to avoid unnecessary consumption of metabolic energy through a futile cycle of cGMP synthesis and breakdown. On the other hand, visual excitation following photic stimulation must generate a very rapid (millisecond) light response that requires immediate acceleration of cGMP breakdown by activated PDE6. Two features that are unique to the PDE6 family of PDEs—the extrinsic regulation of activity by the inhibitory γ-subunit and the extraordinary catalytic efficiency of the PDE6 active site—explain how this member of the PDE superfamily serves as the central effector of visual transduction. In dark-adapted rod outer segments, electrophysiological measurements of “dark noise” suggest that only 1 of 5,000 PDE6 molecules is spontaneously active at any moment [152]. This observation is consistent with measurements of the amount [73] and binding affinity of the γ-subunit for the PDE6 catalytic dimer [86, 142, 153], which together serve to ensure that nonactivated PDE6 exists as a nonactivated tetramer (αβγ2). Direct measurements of the basal activity of PDE6 in rod outer segment suspensions indicate that only 1 of 2,200 PDE6 molecules is spontaneously active in the darkadapted state [154]. On light activation, displacement of the γ-subunit by activated transducin relieves the inhibitory constraints on the PDE6 holoenzyme and triggers the rapid decline in cGMP levels (discussed in detail in next section). Full removal of the γ-subunit from the catalytic dimer by various means (proteolysis [85]; polycationic proteins [78]; or γ-subunit extraction in a complex with activated transducin [155]) results in a PDE6 catalytic dimer that hydrolyzes cGMP with about 1,000-fold greater catalytic efficiency than the closely related PDE5 enzyme (Table 1). The kcat/KM value of 4 × 108 M−1s−1 for catalysis of cGMP by bovine rod PDE6 [142] or amphibian rod PDE6 [154] approaches the diffusion-controlled limit for a bimolecular collision [156], making activated PDE6 a nearly perfectly designed catalyst for degrading cGMP in photoreceptor outer segments during light stimulation [51]. cAMP is a very poor substrate for PDE6 catalysis; the specificity constants (kcat/KM) for cGMP and cAMP differ by about 100-fold, primarily due to differences in the KM values (Table 1). Wide variations in the literature for the KM for PDE6 [40] are now understood to have resulted from restricted substrate diffusion in those experiments when PDE6 activity was measured in rod outer segment membrane preparations [52, 157]. Table 1.

Catalytic properties of human PDE5 and bovine rod PDE6 catalytic dimer Substrate KM (µM) kcat (s−1) kcat/KM (M−1s−1) Reference 8 Rod PDE6 cGMP 14 5440 3.9 × 10 142 cAMP 910 3060 3.4 × 106 PDE5 cGMP 2.9 2.2 7.6 × 105 194 cAMP 290 1.6 5.5 × 103 cAMP cyclic adenosine monophosphate, cGMP cyclic guanosine monophosphate, PDE phosphodiesterase

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PDE6 REGULATION The extent of PDE6 activation and the lifetime of activated PDE6 following photic stimulation are the rate-limiting steps in the excitation and recovery phases of the phototransduction pathway. While binding of activated transducin (specifically the α-subunit with GTP bound; α*-GTP) to relieve the inhibitory constraint of the γ-subunit is central to the PDE6 activation/inactivation mechanism, other factors (such as allosteric regulation by the GAF domains and binding of other PDE6 regulatory proteins) are likely to modulate the light sensitivity, extent of amplification, and duration of the activated state of PDE6. Transducin Activation of Rod PDE6 During Visual Excitation It is generally agreed that transducin activation of PDE6 results from the binding of the activated transducin α-subunit (α*-GTP) to the nonactivated PDE6 holoenzyme (αβγ2) and the displacement of the γ-subunit from its binding site at the entrance to the PDE6 catalytic site (Fig. 4). However, the detailed mechanism of this process remains surprisingly unclear. The prevailing model of visual excitation (Fig. 4A) asserts that one molecule of α*GTP binds to each PDE6 catalytic subunit, displacing both γ-subunits and enhancing cGMP hydrolysis at each catalytic site [45, 158]. However, evidence for the stoichiometry of αt-GTP binding to PDE6 and the extent to which PDE6 can be activated is not consistent with this model. For example, in those instances when physical removal of the γ-subunit from the PDE6 holoenzyme was directly compared to the maximal extent of transducin activation of PDE6, the hydrolytic activity was up to twofold higher for the PDE6 αβ catalytic dimer (devoid of γ-subunit) compared to the α*-GTP–PDE6 activated complex [73, 159–161]. Furthermore, correlations of α*-GTP binding to PDE6 with activation of cGMP hydrolysis demonstrate that a single α*-GTP was able to maximally activate the PDE6 αβ catalytic dimer [161–163]. These observations are consistent with the idea that α*-GTP relieves inhibition at only one of the two active sites on the PDE6 catalytic dimer (Fig. 4B). Although evidence for a second α*-GTP binding to PDE6 (Fig. 4C) has been reported [164], its affinity is likely much weaker and nonproductive under physiological conditions. A description of the mechanism of PDE6 reinhibition following transducin deactivation was presented in this chapter in the sections on deactivation of transducin and PDE6. Functions of the Regulatory cGMP-Binding GAF Domains of PDE6 Whereas allosteric regulation of catalysis has been demonstrated for the PDE2 and PDE5 cGMP-binding GAF domains [165–169], no evidence for intramolecular allosteric communication between the GAF and catalytic domains has been reported for PDE6 [142, 154, 170]. There are, however, inherent experimental difficulties in quantifying cGMP binding to the GAF domains when cGMP is itself a substrate for catalysis at the active site. Nonetheless, cGMP binding to the PDE6 GAF domains must induce a conformational change in the catalytic subunits since the affinity with which the γ-subunit binds to the catalytic dimer is markedly enhanced when cGMP is bound (Fig. 5). This can be seen as a decrease in the basal activity of PDE6 holoenzyme when cGMP occupies

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Fig. 4. Alternative models for the activation of phosphodiesterase 6 (PDE6) by transducin. In model A, a transducin α*-GTP (α-subunit with GTP bound) binds to each of the PDE6 catalytic subunits to displace its bound γ-subunit, leading to full catalytic activity at both active sites. In model B, a single α*-GTP binds to one of the PDE6 catalytic subunits, activating only one of the two catalytic sites in the catalytic dimer. In model C, two α*-GTP bind to PDE6, but only one of the catalytic sites is activated, the other site remaining inhibited.

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Fig. 5. Positive cooperativity between cyclic guanosine monophosphate (cGMP) and γsubunit binding to the rod phosphodiesterase 6 (PDE6) catalytic dimer. The enzyme active sites of PDE6 are denoted by notches, while the cGMP-binding sites are represented by circular pockets. A When γ-subunit (shown with its catalytic-interacting and GAF-interacting subdomains) is mixed with PDE6 catalytic dimer lacking bound cGMP at the GAF domains, the binding affinity of both γ-subunits is equal (dissociation constant, KD = 3 pM). On occupancy of the GAF domains by cGMP (black circles), one γ-subunit-binding site now binds to the catalytic subunit with more than tenfold higher affinity, while the other is unchanged. B Addition of cGMP to the catalytic dimer results in high-affinity (KD = 60 nM) binding to one GAF domain. Only at high cGMP concentrations will the second site become occupied (KD > 1 µM). On addition of stoichiometric amounts of γ-subunit, the cGMP-binding affinity is greatly increased at the low-affinity site and modestly increased at the high-affinity site. (KD values are for bovine rod PDE6 [96, 142].)

the GAF domain [154]. Furthermore, when cGMP is bound to the PDE6 catalytic dimer, the intrinsic γ-subunit binding affinity is enhanced for one, but not both, of its binding sites on the catalytic dimer; the second γ-subunit binding site retains the same affinity for PDE6 regardless of the state of occupancy of the GAF domains by cGMP [142]. When transducin activates the PDE6 holoenzyme, the γ-subunit remains associated with the PDE6 catalytic dimer when cGMP is present but is released in a complex with transducin α-subunit when the GAF domains are unoccupied [170, 171]. This allosteric change in the GAFa domain is reciprocal in that addition of γ-subunit to PDE6 catalytic dimers greatly enhances the binding affinity of cGMP for PDE6 [95, 143]. The two cGMP-binding sites have intrinsically different binding affinities: One GAF domain binds cGMP with high affinity, while the other GAF domain is a low-affinity site in the absence of γ-subunit (Fig. 5B). The cGMP-binding affinity at both sites is increased more than 100-fold when γ-subunit recombines with the catalytic dimer [73, 96, 142]. The reciprocal positive cooperativity between cGMP and γ-subunit binding to PDE6 catalytic dimer is also relevant to transducin-activated PDE6. Not only does displacement of the γ-subunit by α*-GTP relieve inhibition at the active site, cGMP-binding

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affinity to one of the GAF domains is lowered about tenfold. Once cGMP has dissociated from the catalytic subunit, the γ-subunit affinity is concomitantly reduced, causing its dissociation from PDE6, presumably in a complex of with α*-GTP. The second GAF domain retains high affinity for cGMP, and the second γ-subunit remains associated with the PDE6 catalytic dimer [73, 95]. Taken together, the interplay between the γ-subunit and the GAF domains suggests a unique physiological role for the PDE6 GAF domains. In dark-adapted photoreceptors, cytoplasmic free cGMP levels are several micromolar, and the cGMP-binding GAF domains would be occupied with cGMP while two γ-subunits block catalysis at the active sites. On light activation of PDE6, displacement of one γ-subunit by transducin will relieve inhibition at the active site and lower cGMP affinity for one binding site. For transient light activation, cGMP dissociation from the GAFa domains is unlikely because recovery of cGMP levels is fast, thereby promoting tight reassociation of the γ-subunit and the return of PDE6 holoenzyme to its dark-adapted state. For prolonged illumination (i.e., during light adaptation) during which cGMP levels remain low, cGMP dissociation from the GAFa domain might occur, lowering the γ-subunit affinity for its PDE6 catalytic subunit and permitting the γ-subunit (complexed with transducin α-subunit and RGS9) to serve as a GTPase accelerating factor [64]. This would increase the rate of transducin inactivation and help restore PDE6 to its nonactivated state. In this way, the GAFa domains of PDE6 might be sensors of cytoplasmic cGMP and respond to sustained decreases in cGMP levels with a negative-feedback mechanism to help restore the ability to detect light stimuli. An alternative hypothesis that the GAF domains buffer cellular cGMP and release it during photoresponse recovery [67, 97] has not been supported by the kinetics of cGMP binding and dissociation with the GAFa domains [68, 73]. Potential PDE6 Regulatory Binding Proteins Two photoreceptor proteins, a glutamic acid-rich protein 2 (GARP2) and a 17-kDa prenyl-binding protein (PrBP/δ; originally referred to as the PDE “δ-subunit”) have been shown to bind to PDE6 [172, 173], but their roles in regulating PDE6 activity or its subcellular localization are currently unknown. Glutamic Acid-Rich Protein 2 GARP2 is a truncated, alternative splice product of the β-subunit of the rod cGMPgated ion channel (CNGB1). GARP2 has a unique eight amino acid C-terminus (compared to the CNGB1 sequence), a high content of proline and glutamate residues, and a natively unfolded structure [173–176]. GARP2 is specifically expressed in rods but not cones [173] and is concentrated at the rims of the outer segment disk membranes [173, 175]. The GARP2 content in rod outer segments is roughly stoichiometric with the PDE6 content [176, 177], making it an attractive candidate as a PDE6 regulatory protein. The few studies of the ability of GARP2 to regulate PDE6 differ in their conclusions. One study reported that addition of recombinant GARP2 was able to deactivate transducin-activated PDE6, but had no effect on nonactivated PDE6 holoenzyme or the catalytic dimer [173]. In contrast, purified, native GARP2 failed to deactivate transducin-activated PDE6 [177], and it is now believed that this inhibitory action of GARP2 on activated PDE6 can be attributed to the fusion tag present on the recombinant

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GARP2 used in the earlier study [32]. Native GARP2 has been shown to reduce the basal activity of the dark-adapted PDE6 holoenzyme by up to 80% [177]. This result suggests that GARP2 may reinforce the inhibitory action of the γ-subunit on nonactivated PDE6, thereby enhancing the signal-to-noise ratio of rod photoreceptors under very dim (e.g., single-photon) illumination conditions. Other roles for GARP2 have been postulated [173, 176, 178], and it remains to be determined what rod-specific role(s) GARP2 plays in regulating PDE6 or other phototransduction proteins during visual signaling. 17-kDa Prenyl-Binding Protein (PDEδ) The 17-kDa prenyl-binding protein (PrBP/δ) was prematurely identified as the PDE6 δ-subunit because it copurified with a soluble fraction (20–30% of the total) of PDE6 from bovine retinal extracts [172, 179]. Unlike most photoreceptor proteins with tissue distribution that is restricted to the retina, PrBP/δ is ubiquitously expressed [179, 180]. PrBP/δ is capable of interacting with numerous proteins, most—but not all of which—are posttranslationally modified with farnesyl or geranylgeranyl groups (for review, see [181]). The functions of PrBP/δ in photoreceptor cells have been addressed by subcellular localization studies that have variously observed PrBP/δ immunoreactivity in the outer segments, the inner segments, as well as the connecting cilium region between inner and outer segments of both rods and cones [179, 182–184]. Localization of PrBP/δ in the inner segment and the connecting cilium region of the photoreceptor is consistent with a role for PrBP/δ in the transport of prenylated proteins from their site of synthesis in the inner segment to the outer segment. This idea is supported by identification of PrBP/ δ-binding partners that are thought to be involved in vesicular transport [185–187]. Recent work with a transgenic mouse in which the PrBP/δ gene has been deleted shows a reduction in transport of opsin kinase (a prenylated protein) to the outer segments of rods and cones, with corresponding defects in the photoresponse [183]. Biochemical studies have shown that PrBP/δ is a high-affinity PDE6-binding protein, with specific interaction with the farnesylated and geranylgeranylated C-termini of rod PDE6 catalytic subunits. Binding of PrBP/δ with PDE6 releases PDE6 from its disk membrane attachment site [179, 184, 188]. Once solubilized, the ability of activated transducin α-subunit to activate PDE6 is greatly impaired [184, 189]. In addition, PrBP/δ binding to PDE6 reduces cGMP-binding affinity to one of the two GAF domain binding sites [96] and disrupts the high-affinity interaction of PDE6 with GARP2 [177]. These specific effects of PrBP/δ on PDE6 in vitro support a role for PrBP/δ in negativefeedback regulation of PDE6 activation, perhaps during prolonged light adaptation. However, the PrBP/δ content in rod outer segments is only 10% of the PDE6 concentration [184], arguing against PrBP/δ as a bona fide PDE6 regulatory subunit. More work is needed to fully understand the role of PrBP/δ in the phototransduction pathway or in protein transport in photoreceptor cells. CONCLUSIONS Considering the wealth of knowledge about the phototransduction pathway—arguably the most thoroughly characterized of the G protein-coupled signaling pathways—it is humbling to realize that major gaps remain in understanding the complexities of PDE6

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regulation at the center of the cGMP signaling cascade. The detailed sequence of events leading to transducin activation of the PDE6 holoenzyme remains to be determined. The physiological significance of the allosteric communication of the PDE6 GAF domains through the inhibitory γ-subunit remains speculative at present. Some aspects of the complex set of processes known as “light adaptation” are postulated to operate by novel mechanisms [190–192]. Novel PDE6 regulatory proteins are possible candidates to modulate PDE6 activity during light adaptation. Finally, this review focused almost exclusively on visual transduction in rod photoreceptors, in large part because so little is known about the extent to which differences between rod and cone PDE6 regulation may account for some of the physiological differences in rod and cone photoresponses [193]. PDE6 is coming under increasing scrutiny outside of the field of vision research. With the increasing reliance on PDE inhibitors for a growing number of therapeutic applications (e.g., male erectile dysfunction, pulmonary hypertension), potential adverse effects on PDE6 function must be considered. There is also limited evidence to date that PDE6 subunits may be found outside the retina and may be differentially expressed during development. Having a G protein-coupled PDE6 expressed in nonphotoreceptive cells is extremely intriguing and merits further investigation of how PDE6 may participate in regulating cyclic nucleotide levels in other cells and tissues. ACKNOWLEDGMENTS Work from my laboratory is supported by the National Eye Institute (EY05798). This is Scientific Contribution 2313 from the New Hampshire Agricultural Experiment Station. REFERENCES 1. Lincoln, T. M. (1994). Cyclic GMP: biochemistry, physiology and pathophysiology. Landes, Austin, TX. 2. Corbin, J. D., Kotera, J., Gopal, V. K., Cote, R. H., Francis, S. H. (2003). Regulation of cyclic nucleotide levels by sequestration. In: Handbook of cell signaling (Bradshaw, R., Dennis, E., eds.), pp. 465–470, Academic Press, San Diego, CA. 3. Sager, G. (2004). Cyclic GMP transporters. Neurochem. Int. 45, 865–873. 4. Pugh, E. N., Jr., Duda, T., Sitaramayya, A., Sharma, R. K. (1997). Photoreceptor guanylate cyclases: a review. Biosci. Rep. 17, 429–473. 5. Kuhn, M. (2003). Structure, regulation, and function of mammalian membrane guanylyl cyclase receptors, with a focus on guanylyl cyclase-A. Circ. Res. 93, 700–709. 6. Olshevskaya, E. V., Ermilov, A. N., Dizhoor, A. M. (2002). Factors that affect regulation of cGMP synthesis in vertebrate photoreceptors and their genetic link to human retinal degeneration. Mol. Cell. Biochem. 230, 139–147. 7. Koch, K. W., Duda, T., Sharma, R. K. (2002). Photoreceptor specific guanylate cyclases in vertebrate phototransduction. Mol. Cell. Biochem. 230, 97–106. 8. Beavo, J. A., Francis, S. H., Houslay, M. D. (2006). Cyclic nucleotide phosphodiesterases in health and disease. CRC Press, Boca Raton, FL. 9. Cote, R. H. (2004). Characteristics of photoreceptor PDE (PDE6): similarities and differences to PDE5. Int. J. Impot. Res. 16, S28–S33. 10. Kruh, G. D., Belinsky, M. G. (2003). The MRP family of drug efflux pumps. Oncogene 22, 7537–7552. 11. Chen, Z. S., Guo, Y., Belinsky, M. G., Kotova, E., Kruh, G. D. (2005). Transport of bile acids, sulfated steroids, estradiol 17-β-D-glucuronide, and leukotriene C4 by human multidrug resistance protein 8 (ABCC11). Mol. Pharmacol. 67, 545–557.

160

Cote

12. Nakatani, K., Yau, K.-W. (1988). Guanosine 3′–5′-cyclic monophosphate-activated conductance studied in a truncated rod outer segment of the toad. J. Physiol. (Lond.) 395, 731–753. 13. Pugh, E. N., Jr., Lamb, T. D. (1990). Cyclic GMP and calcium: the internal messengers of excitation and adaptation in vertebrate photoreceptors. Vision Res. 30, 1923–1948. 14. Cote, R. H., Biernbaum, M. S., Nicol, G. D., Bownds, M. D. (1984). Light-induced decreases in cGMP concentration precede changes in membrane permeability in frog rod photoreceptors. J. Biol. Chem. 259, 9635–9641. 15. Hamm, H. (1990). Regulation by light of cyclic nucleotide-dependent protein kinases and their substrates in frog rod outer segments. J. Gen. Physiol. 95, 545–567. 16. Gamm, D. M., Barthel, L. K., Raymond, P. A., Uhler, M. D. (2000). Localization of cGMPdependent protein kinase isoforms in mouse eye. Invest. Ophthalmol. Vis. Sci. 41, 2766– 2773. 17. Lee, R. H., Brown, B. M., Lolley, R. N. (1981). Protein kinases of retinal rod outer segments: identification and partial characterization of cyclic nucleotide dependent protein kinase and rhodopsin kinase. Biochemistry 20, 7532–7538. 18. Binder, B. M., Brewer, E., Bownds, M. D. (1989). Stimulation of protein phosphorylations in frog rod outer segments by protein kinase activators. J. Biol. Chem. 264, 8857–8864. 19. Polans, A. S., Hermolin, J., Bownds, M. D. (1979). Light-induced dephosphorylation of two proteins in frog rod outer segments: influence of cyclic nucleotides and calcium. J. Gen. Physiol. 74, 595–613. 20. Hermolin, J., Karell, M. A., Hamm, H. E., Bownds, M. D. (1982). Calcium and cyclic GMP regulation of light-sensitive protein phosphorylation in frog photoreceptor membranes. J. Gen. Physiol. 79, 633–655. 21. Lee, R. H., Brown, B. M., Lolley, R. N. (1984). Light-induced dephosphorylation of a 33 K protein in rod outer segments of rat retina. Biochemistry 23, 1972–1977. 22. Shinozawa, T., Yoshizawa, T. (1986). Cyclic nucleotide-dependent phosphorylation of proteins in rod outer segments in frog retina. J. Biol. Chem. 261, 216–223. 23. Krapivinskii, G. B., Malenev, A. L., Fesenko, E. E. (1987). Cyclonucleotide- and lightdependent phosphorylation of rod outer segment proteins from frog retina. Mol. Biol. (Mosc.) 21, 116–124. 24. Pagh-Roehl, K., Han, E., Burnside, B. (1993). Identification of cyclic nucleotide-regulated phosphoproteins, including phosducin, in motile rod inner-outer segments of teleosts. Exp. Eye Res. 57, 679–691. 25. Suh, K. H., Hamm, H. E. (1996). Cyclic AMP-dependent phosphoprotein components I and II interact with βgamma subunits of transducin in frog rod outer segments. Biochemistry 35, 290–298. 26. Bauer, P. H., Muller, S., Puzicha, M., Pippig, S., Obermaier, B., Helmreich, E. J. M., Lohse, M. J. (1992). Phosducin is a protein kinase A-regulated G-protein regulator. Nature 358, 73–76. 27. Gaudet, R., Savage, J. R., McLaughlin, J. N., Willardson, B. M., Sigler, P. B. (1999). A molecular mechanism for the phosphorylation-dependent regulation of heterotrimeric G proteins by phosducin. Mol. Cell 3, 649–660. 28. Sokolov, M., Strissel, K. J., Leskov, I. B., Michaud, N. A., Govardovskii, V. I., Arshavsky, V. Y. (2004). Phosducin facilitates light-driven transducin translocation in rod photoreceptors: Evidence from the phosducin knockout mouse. J. Biol. Chem. 279, 19149–19156. 29. Matulef, K., Zagotta, W. N. (2003). Cyclic nucleotide-gated ion channels. Annu. Rev. Cell Dev. Biol. 19, 23–44. 30. Zimmerman, A. L. (2002). Two B or not two B? Questioning the rotational symmetry of tetrameric ion channels. Neuron 36, 997–999. 31. Peng, C., Rich, E. D., Varnum, M. D. (2004). Subunit configuration of heteromeric cone cyclic nucleotide-gated channels. Neuron 42, 401–410.

cGMP Signaling Pathway and Role of PDE6

161

32. Kaupp, U. B., Seifert, R. (2002). Cyclic nucleotide-gated ion channels. Physiol. Rev. 82, 769–824. 33. Molday, R. S., Kaupp, U. B. (2000). Ion channels of vertebrate photoreceptors. In: Molecular mechanisms in visual transduction (Stavenga, D. G., DeGrip, W. J., Pugh, E. N., Jr., eds.), pp. 143–181, North Holland, Amsterdam. 34. Kramer, R. H., Molokanova, E. (2001). Modulation of cyclic-nucleotide-gated channels and regulation of vertebrate phototransduction. J. Exp. Biol. 204, 2921–2931. 35. McCabe, S. L., Pelosi, D. M., Tetreault, M., Miri, A., Nguitragool, W., Kovithvathanaphong, P., Mahajan, R., Zimmerman, A. L. (2004). All-trans-retinal is a closed-state inhibitor of rod cyclic nucleotide-gated ion channels. J. Gen. Physiol. 123, 521–531. 36. Bradley, J., Reisert, J., Frings, S. (2005). Regulation of cyclic nucleotide-gated channels. Curr. Opin. Neurobiol. 15:343–349 37. Rodieck, R. W. (1998). The first steps in seeing. Sinauer Associates, Sunderland, MA. 38. Fain, G. L. (2003). Sensory transduction. Sinauer, Sunderland, MA. 39. Baylor, D. A. (1987). Photoreceptor signals and vision. Invest. Ophthalmol. Vis. Sci. 28, 34– 49. 40. Pugh, E. N., Jr., Lamb, T. D. (1993). Amplification and kinetics of the activation steps in phototransduction. Biochim. Biophys. Acta 1141, 111–149. 41. Yau, K.-W. (1994). Phototransduction mechanism in retinal rods and cones. Invest. Ophthalmol. Vis. Sci. 35, 9–32. 42. Rieke, F., Baylor, D. A. (1998). Single photon detection by rod cells of the retina. Rev. Mod. Phys. 70, 1027–1036. 43. Nikonov, S., Engheta, H., Pugh, E. N., Jr. (1998). Kinetics of recovery of the dark-adapted salamander rod photoresponse. J. Gen. Physiol. 111, 7–37. 44. Pugh, E. N., Jr., Nikonov, S., Lamb, T. D. (1999). Molecular mechanisms of vertebrate photoreceptor light adaptation. Curr. Opin. Neurobiol. 9, 410–418. 45. Pugh, E. N., Lamb, T. D. (2000). Phototransduction in vertebrate rods and cones: molecular mechanisms of amplification, recovery and light adaptation. In: Molecular mechanisms in visual transduction (Stavenga, D. G., DeGrip, W. J., Pugh, E. N., eds.), pp. 183–255, Elsevier Science, New York. 46. Fain, G. L., Matthews, H. R., Cornwall, M. C., Koutalos, Y. (2001). Adaptation in vertebrate photoreceptors. Physiol. Rev. 81, 117–151. 47. Pugh, E. N. (2006). RGS expression level precisely regulates the duration of rod photoresponses. Neuron 51, 391–393. 48. Filipek, S., Stenkamp, R. E., Teller, D. C., Palczewski, K. (2003). G protein-coupled receptor rhodopsin: a prospectus. Annu. Rev. Physiol. 65, 26.1–26.29. 49. Hubbell, W. L., Altenbach, C., Hubbell, C. M., Khorana, H. G. (2003). Rhodopsin structure, dynamics, and activation: a perspective from crystallography, site-directed spin labeling, sulfhydryl reactivity, and disulfide cross-linking. Adv. Protein Chem. 63, 243–290. 50. Cabrera-Vera, T. M., Vanhauwe, J., Thomas, T. O., Medkova, M., Preininger, A., Mazzoni, M. R., Hamm, H. E. (2003). Insights into G protein structure, function, and regulation. Endocr. Rev. 24, 765–781. 51. Arshavsky, V. Y., Lamb, T. D., Pugh, E. N., Jr. (2002). G proteins and phototransduction. Annu. Rev. Physiol. 64, 153–187. 52. Leskov, I. B., Klenchin, V. A., Handy, J. W., Whitlock, G. G., Govardovskii, V. I., Bownds, M. D., Lamb, T. D., Pugh, E. N., Jr., Arshavsky, V. Y. (2000). The gain of rod phototransduction: Reconciliation of biochemical and electrophysiological measurements. Neuron 27, 525–537. 53. Krispel, C. M., Chen, D., Melling, N., Chen, Y. J., Martemyanov, K. A., Quillinan, N., Arshavsky, V. Y., Wensel, T. G., Chen, C. K., Burns, M. E. (2006). RGS expression ratelimits recovery of rod photoresponses. Neuron 51, 409–416.

162

Cote

54. Burns, M. E., Arshavsky, V. Y. (2005). Beyond counting photons: trials and trends in vertebrate visual transduction. Neuron 48, 387–401. 55. Kawamura, S., Tachibanaki, S. (2002). S-modulin. Adv. Exp. Med. Biol. 514, 61–68. 56. Senin, I. I., Koch, K. W., Akhtar, M., Philippov, P. P. (2002). Ca2+-dependent control of rhodopsin phosphorylation: recoverin and rhodopsin kinase. Adv. Exp. Med. Biol. 514, 69–99. 57. Maeda, T., Imanishi, Y., Palczewski, K. (2003). Rhodopsin phosphorylation: 30 years later. Progr. Retin. Eye Res. 22, 417–434. 58. Makino, C. L., Dodd, R. L., Chen, J., Burns, M. E., Roca, A., Simon, M. I., Baylor, D. A. (2004). Recoverin regulates light-dependent phosphodiesterase activity in retinal rods. J. Gen. Physiol. 123, 729–741. 59. Gurevich, V. V., Gurevich, E. V. (2004). The molecular acrobatics of arrestin activation. Trends Pharmacol. Sci. 25, 105–111. 60. Lamb, T. D., Pugh, E. N. (2004). Dark adaptation and the retinoid cycle of vision. Prog. Retin. Eye Res. 23, 307–380. 61. He, W., Cowan, C. W., Wensel, T. G. (1998). RGS9, a GTPase accelerator for phototransduction. Neuron 20, 95–102. 62. Makino, E. R., Handy, J. W., Li, T. S., Arshavsky, V. Y. (1999). The GTPase activating factor for transducin in rod photoreceptors is the complex between RGS9 and type 5 G protein β subunit. Proc. Natl. Acad. Sci. U. S. A. 96, 1947–1952. 63. Hu, G., Wensel, T. G. (2002). R9AP, a membrane anchor for the photoreceptor GTPase accelerating protein, RGS9-1. Proc. Natl. Acad. Sci. U. S. A. 99, 9755–9760. 64. Arshavsky, V. Y., Bownds, M. D. (1992). Regulation of deactivation of photoreceptor G protein by its target enzyme and cGMP. Nature 357, 416–417. 65. Skiba, N. P., Hopp, J. A., Arshavsky, V. Y. (2000). The effector enzyme regulates the duration of G protein signaling in vertebrate photoreceptors by increasing the affinity between transducin and RGS protein. J. Biol. Chem. 275, 32716–32720. 66. Slep, K. C., Kercher, M. A., He, W., Cowan, C. W., Wensel, T. G., Sigler, P. B. (2001). Structural determinants for regulation of phosphodiesterase by a G protein at 2.0 A. Nature 409, 1071–1077. 67. Yamazaki, A., Bondarenko, V. A., Dua, S., Yamazaki, M., Usukura, J., Hayashi, F. (1996). Possible stimulation of retinal rod recovery to dark state by cGMP release from a cGMP phosphodiesterase noncatalytic site. J. Biol. Chem. 271, 32495–32498. 68. Calvert, P. D., Ho, T. W., LeFebvre, Y. M., Arshavsky, V. Y. (1998). Onset of feedback reactions underlying vertebrate rod photoreceptor light adaptation. J. Gen. Physiol. 111, 39–51. 69. Erickson, M. A., Robinson, P., Lisman, J. (1992). Deactivation of visual transduction without guanosine triphosphate hydrolysis by G protein. Science 257, 1255–1258. 70. Yamazaki, A., Moskvin, O., Yamazaki, R. K. (2002). Phosphorylation by cyclin-dependent protein kinase 5 of the regulatory subunit (Pgamma) of retinal cgmp phosphodiesterase (PDE6): its implications in phototransduction. Adv. Exp. Med. Biol. 514, 131–153. 71. Tsang, S. H., Woodruff, M. L., Chen, C. K., Yamashita, C. K., Cilluffo, M. C., Rao, A. L., Farber, D. B., Fain, G. L. (2006). GAP-independent termination of photoreceptor light response by excess γ subunit of the cGMP-phosphodiesterase. J. Neurosci. 26, 4472–4480. 72. Sitaramayya, A., Harkness, J., Parkes, J. H., Gonzalez-Oliva, C., Liebman, P. A. (1986). Kinetic studies suggest that light-activated cyclic GMP phosphodiesterase is a complex with G-protein subunits. Biochemistry 25, 651–656. 73. Norton, A. W., D’Amours, M. R., Grazio, H. J., Hebert, T. L., Cote, R. H. (2000). Mechanism of transducin activation of frog rod photoreceptor phosphodiesterase: allosteric interactions between the inhibitory γ subunit and the noncatalytic cGMP binding sites. J. Biol. Chem. 275, 38611–38619.

cGMP Signaling Pathway and Role of PDE6

163

74. Hsu, Y.-T., Molday, R. S. (1993). Modulation of the cGMP-gated channel of rod photoreceptor cells by calmodulin. Nature 361, 76–79. 75. Korenbrot, J. I., Rebrik, T. I. (2002). Tuning outer segment Ca2+ homeostasis to phototransduction in rods and cones. Adv. Exp. Med. Biol. 514, 179–203. 76. Rebrik, T. I., Korenbrot, J. I. (2004). In intact mammalian photoreceptors, Ca2+-dependent modulation of cGMP-gated ion channels is detectable in cones but not in rods. J. Gen. Physiol. 123, 63–76. 77. Zoraghi, R., Corbin, J. D., Francis, S. H. (2004). Properties and functions of GAF domains in cyclic nucleotide phosphodiesterases and other proteins. Mol. Pharmacol. 65, 267–278. 78. Miki, N., Baraban, J. M., Keirns, J. J., Boyce, J. J., Bitensky, M. W. (1975). Purification and properties of the light-activated cyclic nucleotide phosphodiesterase of rod outer segments. J. Biol. Chem. 250, 6320–6327. 79. Baehr, W., Devlin, M. J., Applebury, M. L. (1979). Isolation and characterization of cGMP phosphodiesterase from bovine rod outer segments. J. Biol. Chem. 254, 11669–11677. 80. Fung, B. K. K., Young, J. H., Yamane, H. K., Griswold-Prenner, I. (1990). Subunit stoichiometry of retinal rod cGMP phosphodiesterase. Biochemistry 29, 2657–2664. 81. Artemyev, N. O., Surendran, R., Lee, J. C., Hamm, H. E. (1996). Subunit structure of rod cGMP-phosphodiesterase. J. Biol. Chem. 271, 25382–25388. 82. Hurwitz, R. L., Bunt Milam, A. H., Chang, M. L., Beavo, J. (1985). cGMP phosphodiesterase in rod and cone outer segments of the retina. J. Biol. Chem. 260, 568–573. 83. Gillespie, P. G., Beavo, J. A. (1988). Characterization of a bovine cone photoreceptor phosphodiesterase purified by cyclic GMP-Sepharose chromatography. J. Biol. Chem. 263, 8133–8141. 84. Dumler, I. L., Etingof, R. N. (1976). Protein inhibitor of cyclic adenosine 3′:5′-monophosphate phosphodiesterase in retina. Biochim. Biophys. Acta 429, 474–478. 85. Hurley, J. B., Stryer, L. (1982). Purification and characterization of the gamma regulatory subunit of the cyclic GMP phosphodiesterase from retinal rod outer segments. J. Biol. Chem. 257, 11094–11099. 86. Deterre, P., Bigay, J., Forquet, F., Robert, M., Chabre, M. (1988). cGMP phosphodiesterase of retinal rods is regulated by two inhibitory subunits. Proc. Natl. Acad. Sci. U. S. A. 85, 2424–2428. 87. Ovchinnikov, Y. A., Gubanov, V. V., Khramtxov, N. V., Ischenko, K. A., Zagranichny, V. E., Muradov, K. G., Shuvaeva, T. M., Lipkin, V. M. (1987). Cyclic GMP phosphodiesterase from bovine retina: amino acid sequence of the alpha-subunit and nucleotide sequence of the corresponding cDNA. FEBS Lett. 223, 169–173. 88. Lipkin, V. M., Khramtsov, N. V., Vasilevskaya, I. A., Atabekova, N. V., Muradov, K. G., Gubanov, V. V., Li, T., Johnston, J. P., Volpp, K. J., Applebury, M. L. (1990). β-Subunit of bovine rod photoreceptor cGMP phosphodiesterase: comparison with the phosphodiesterase family. J. Biol. Chem. 265, 12955–12959. 89. Li, T., Volpp, K., Applebury, M. L. (1990). Bovine cone photoreceptor cGMP phosphodiesterase structure deduced from a cDNA clone. Proc. Natl. Acad. Sci. U. S. A. 87, 293–297. 90. Aravind, L., Ponting, C. P. (1997). The GAF domain: an evolutionary link between diverse phototransducing proteins. Trends Biochem. Sci. 22, 458–459. 91. Yamazaki, A., Sen, I., Bitensky, M. W., Casnellie, J. E., Greengard, P. (1980). Cyclic GMPspecific, high affinity, noncatalytic binding sites on light-activated phosphodiesterase. J. Biol. Chem. 255, 11619–11624. 92. Muradov, K. G., Boyd, K. K., Martinez, S. E., Beavo, J. A., Artemyev, N. O. (2003). The GAFa domains of rod cGMP-phosphodiesterase 6 determine the selectivity of the enzyme dimerization. J. Biol. Chem. 278, 10594–10601.

164

Cote

93. Muradov, K. G., Boyd, K. K., Artemyev, N. O. (2005). Analysis of dimerization determinants of PDE6 catalytic subunits. Methods Mol. Biol. 307, 263–276. 94. Gillespie, P. G., Beavo, J. A. (1989). cGMP is tightly bound to bovine retinal rod phosphodiesterase. Proc. Natl. Acad. Sci. U. S. A. 86, 4311–4315. 95. Cote, R. H., Bownds, M. D., Arshavsky, V. Y. (1994). cGMP binding sites on photoreceptor phosphodiesterase: role in feedback regulation of visual transduction. Proc. Natl. Acad. Sci. U. S. A. 91, 4845–4849. 96. Mou, H., Grazio, H. J., Cook, T. A., Beavo, J. A., Cote, R. H. (1999). cGMP binding to noncatalytic sites on mammalian rod photoreceptor phosphodiesterase is regulated by binding of its γ and δ subunits. J. Biol. Chem. 274, 18813–18820. 97. Cote, R. H., Brunnock, M. A. (1993). Intracellular cGMP concentration in rod photoreceptors is regulated by binding to high and moderate affinity cGMP binding sites. J. Biol. Chem. 268, 17190–17198. 98. Hebert, M. C., Schwede, F., Jastorff, B., Cote, R. H. (1998). Structural features of the noncatalytic cGMP binding sites of frog photoreceptor phosphodiesterase using cGMP analogs. J. Biol. Chem. 273, 5557–5565. 99. Huang, D., Hinds, T. R., Martinez, S. E., Doneanu, C., Beavo, J. A. (2004). Molecular determinants of cGMP-binding to chicken cone photoreceptor phosphodiesterase. J. Biol. Chem. 279, 48143–48151. 100. Muradov, H., Boyd, K. K., Artemyev, N. O. (2004). Structural determinants of the PDE6 GAF A domain for binding the inhibitory gamma-subunit and noncatalytic cGMP. Vision Res. 44, 2437–2444. 101. Muradov, K. G., Granovsky, A. E., Schey, K. L., Artemyev, N. O. (2002). Direct interaction of the inhibitory γ-subunit of rod cGMP phosphodiesterase (PDE6) with the PDE6 GAFa domains. Biochemistry 41, 3884–3890. 102. Guo, L. W., Grant, J. E., Hajipour, A. R., Muradov, H., Arbabian, M., Artemyev, N. O., Ruoho, A. E. (2005). Asymmetric interaction between rod cyclic GMP phosphodiesterase γ subunits and αβ subunits. J. Biol. Chem. 280, 12585–12592. 103. Kameni Tcheudji, J. F., Lebeau, L., Virmaux, N., Maftei, C. G., Cote, R. H., Lugnier, C., Schultz, P. (2001). Molecular organization of bovine rod cGMP-phosphodiesterase 6. J. Mol. Biol. 310, 781–791. 104. Qin, N., Baehr, W. (1994). Expression and mutagenesis of mouse rod photoreceptor cGMP phosphodiesterase. J. Biol. Chem. 269, 3265–3271. 105. Granovsky, A. E., Natochin, M., McEntaffer, R. L., Haik, T. L., Francis, S. H., Corbin, J. D., Artemyev, N. O. (1998). Probing domain functions of chimeric PDE6α′/PDE5 cGMPphosphodiesterase. J. Biol. Chem. 273, 24485–24490. 106. Piriev, N. I., Yamashita, C. K., Shih, J., Farber, D. B. (2003). Expression of cone photoreceptor cGMP-phosphodiesterase alpha′ subunit in Chinese hamster ovary, 293 human embryonic kidney, and Y79 retinoblastoma cells. Mol. Vis. 9, 80–86. 107. Zhang, J., Kuvelkar, R., Wu, P., Egan, R. W., Billah, M. M., Wang, P. (2004). Differential inhibitor sensitivity between human recombinant and native photoreceptor cGMPphosphodiesterases (PDE6s). Biochem. Pharmacol. 68, 867–873. 108. Muradov, H., Boyd, K. K., Artemyev, N. O. (2006). Analysis of PDE6 function using chimeric PDE5/6 catalytic domains. Vision Res. 46, 860–868. 109. Granovsky, A. E., Artemyev, N. O. (2001). Partial reconstitution of photoreceptor cGMP phosphodiesterase characteristics in cGMP phosphodiesterase-5. J. Biol. Chem. 276, 21698–21703. 110. Francis, S. H., Turko, I. V., Grimes, K. A., Corbin, J. D. (2000). Histidine-607 and histidine-643 provide important interactions for metal support of catalysis in phosphodiesterase-5. Biochemistry 39, 9591–9596.

cGMP Signaling Pathway and Role of PDE6

165

111. Francis, S. H., Colbran, J. L., McAllister-Lucas, L. M., Corbin, J. D. (1994). Zinc interactions and conserved motifs of the cGMP-binding cGMP-specific phosphodiesterase suggest that it is a zinc hydrolase. J. Biol. Chem. 269, 22477–22480. 112. He, F., Seryshev, A. B., Cowan, C. W., Wensel, T. G. (2000). Multiple zinc binding sites in retinal rod cGMP phosphodiesterase, PDE6αβ. J. Biol. Chem. 275, 20572–20577. 113. Artemyev, N. O., Natochin, M., Busman, M., Schey, K. L., Hamm, H. E. (1996). Mechanism of photoreceptor cGMP phosphodiesterase inhibition by its gamma-subunits. Proc. Natl. Acad. Sci. U. S. A. 93, 5407–5412. 114. Granovsky, A. E., Natochin, M., Artemyev, N. O. (1997). The γ subunit of rod cGMPphosphodiesterase blocks the enzyme catalytic site. J. Biol. Chem. 272, 11686–11689. 115. Granovsky, A. E., Artemyev, N. O. (2000). Identification of the γ-subunit interacting residues on photoreceptor cGMP phosphodiesterase, PDE6α′. J. Biol. Chem. 275, 41258–41262. 116. Qin, N., Pittler, S. J., Baehr, W. (1992). In vitro isoprenylation and membrane association of mouse rod photoreceptor cGMP phosphodiesterase α-and β-subunits expressed in bacteria. J. Biol. Chem. 267, 8458–8463. 117. Anant, J. S., Ong, O. C., Xie, H., Clarke, S., O’Brien, P. J., Fung, B. K. K. (1992). In vivo differential prenylation of retinal cyclic GMP phosphodiesterase catalytic subunits. J. Biol. Chem. 267, 687–690. 118. Catty, P., Deterre, P. (1991). Activation and solubilization of the retinal cGMP-specific phosphodiesterase by limited proteolysis—role of the C- terminal domain of the β-subunit. Eur. J. Biochem. 199, 263–269. 119. Kuhn, H. (1980). Light- and GTP-regulated interaction of GTPase and other proteins with bovine photoreceptor membranes. Nature 283, 587–589. 120. Ovchinnikov, Y. A., Lipkin, V. M., Kimarev, V. P., Gubanov, V. V., Khramtsov, N. V., Akhmedov, N. B., Zagranichny, V. E., Muradov, K. G. (1986). Cyclic GMP phosphodiesterase from cattle retina: Amino acid sequence of the γ-subunit and nucleotide sequence of the corresponding cDNA. FEBS Lett. 204, 288–292. 121. Hamilton, S. E., Hurley, J. B. (1990). A phosphodiesterase inhibitor specific to a subset of bovine retinal cones. J. Biol. Chem. 265, 11259–11264. 122. Berger, A. L., Cerione, R. A., Erickson, J. W. (1997). Real time conformation changes in the retinal phosphodiesterase gamma subunit monitored by resonance energy transfer. J. Biol. Chem. 272, 2714–2721. 123. Uversky, V. N., Permyakov, S. E., Zagranichny, V. E., Rodionov, I. L., Fink, A. L., Cherskaya, A. M., Wasserman, L. A., Permyakov, E. A. (2002). Effect of zinc and temperature on the conformation of the gamma subunit of retinal phosphodiesterase: a natively unfolded protein. J. Proteome Res. 1, 149–159. 124. Morin, F., Vannier, B., Houdart, F., Regnacq, M., Berges, T., Voisin, P. (2003). A prolinerich domain in the gamma subunit of phosphodiesterase 6 mediates interaction with SH3containing proteins. Mol. Vis. 9, 449–459. 125. Tsuboi, S., Matsumoto, H., Jackson, K. W., Tsujimoto, K., Williams, T., Yamazaki, A. (1994). Phosphorylation of an inhibitory subunit of cGMP phosphodiesterase in Rana catesbiana rod photoreceptors. I. Characterization of the phosphorylation. J. Biol. Chem. 269, 15016–15023. 126. Matsuura, I., Bondarenko, V. A., Maeda, T., Kachi, S., Yamazaki, M., Usukura, J., Hayashi, F., Yamazaki, A. (2000). Phosphorylation by cyclin-dependent protein kinase 5 of the regulatory subunit of retinal cGMP phosphodiesterase: I. Identification of the kinase and its role in the turnoff of phosphodiesterase in vitro. J. Biol. Chem. 275, 32950–32957. 127. Paglia, M. J., Mou, H., Cote, R. H. (2002). Regulation of photoreceptor phosphodiesterase (PDE6) by phosphorylation of its inhibitory γ subunit re-evaluated. J. Biol. Chem. 277, 5017–5023.

166

Cote

128. Lipkin, V. M., Dumler, I. L., Muradov, K. G., Artemyev, N. O., Etingof, R. N. (1988). Active sites of the cyclic GMP phosphodiesterase gamma-subunit of retinal rod outer segments. FEBS Lett. 234, 287–290. 129. Morrison, D. F., Cunnick, J. M., Oppert, B., Takemoto, D. J. (1989). Interaction of the gamma subunit of retinal rod outer segment phosphodiesterase with transducin. J. Biol. Chem. 264, 11671–11681. 130. Artemyev, N. O., Rarick, H. M., Mills, J. S., Skiba, N. P., Hamm, H. E. (1992). Sites of interaction between rod G-protein α-subunit and cGMP-phosphodiesterase gammasubunit. Implications for phosphodiesterase activation mechanism. J. Biol. Chem. 267, 25067–25072. 131. Brown, R. L. (1992). Functional regions of the inhibitory subunit of retinal rod cGMP phosphodiesterase identified by site-specific mutagenesis and fluorescence spectroscopy. Biochemistry 31, 5918–5925. 132. Artemyev, N. O., Mills, J. S., Thornburg, K. R., Knapp, D. R., Schey, K. L., Hamm, H. E. (1993). A site on transducin α-subunit of interaction with the polycationic region of cGMP phosphodiesterase inhibitory subunit. J. Biol. Chem. 268, 23611–23615. 133. Skiba, N. P., Bae, H., Hamm, H. E. (1996). Mapping of effector binding sites of transducin α-subunit using Gαt/Gαi1 chimeras. J. Biol. Chem. 271, 413–424. 134. Udovichenko, I. P., Cunnick, J., Gonzales, K., Takemoto, D. J. (1993). Phosphorylation of bovine rod photoreceptor cyclic GMP phosphodiesterase. Biochem. J. 295, 49–55. 135. Xu, L. X., Tanaka, Y., Bonderenko, V. A., Matsuura, I., Matsumoto, H., Yamazaki, A., Hayashi, F. (1998). Phosphorylation of the gamma subunit of the retinal photoreceptor cGMP phosphodiesterase by the cAMP-dependent protein kinase and its effect on the gamma subunit interaction with other proteins. Biochemistry 37, 6205–6213. 136. Bondarenko, V. A., Desai, M., Dua, S., Yamazaki, M., Amin, R. H., Yousif, K. K., Kinumi, T., Ohashi, M., Komori, N., Matsumoto, H., Jackson, K. W., Hayashi, F., Usukura, J., Lipkin, V. M., Yamazaki, A. (1997). Residues within the polycationic region of cGMP phosphodiesterase gamma subunit crucial for the interaction with transducin α subunit—identification by endogenous ADP-ribosylation and site-directed mutagenesis. J. Biol. Chem. 272, 15856–15864. 137. Bondarenko, V. A., Yamazaki, M., Hayashi, F., Yamazaki, A. (1999). Suppression of GTP/ Tα-dependent activation of cGMP phosphodiesterase by ADP-ribosylation by its gamma subunit in amphibian rod photoreceptor membranes. Biochemistry 38, 7755–7763. 138. Artemyev, N. O., Hamm, H. E. (1992). Two-site high-affinity interaction between inhibitory and catalytic subunits of rod cyclic GMP phosphodiesterase. Biochem. J. 283, 273–279. 139. Takemoto, D. J., Hurt, D., Oppert, B., Cunnick, J. (1992). Domain mapping of the retinal cyclic GMP phosphodiesterase gamma-subunit. Function of the domains encoded by the three exons of the gamma-subunit gene. Biochem. J. 281, 637–643. 140. Lipkin, V. M., Bondarenko, V. A., Zagranichny, V. E., Dobrynina, L. N., Muradov, K. G., Natochin, M. Y. (1993). Site-directed mutagenesis of the cGMP phosphodiesterase gamma subunit from bovine rod outer segments: Role of separate amino acid residues in the interaction with catalytic subunits and transducin α subunit. Biochim. Biophys. Acta 1176, 250–256. 141. Natochin, M., Artemyev, N. O. (1996). An interface of interaction between photoreceptor cGMP phosphodiesterase catalytic subunits and inhibitory gamma subunits. J. Biol. Chem. 271, 19964–19969. 142. Mou, H., Cote, R. H. (2001). The catalytic and GAF domains of the rod cGMP phosphodiesterase (PDE6) heterodimer are regulated by distinct regions of its inhibitory γ subunit. J. Biol. Chem. 276, 27527–27534.

cGMP Signaling Pathway and Role of PDE6

167

143. Yamazaki, A., Bartucci, F., Ting, A., Bitensky, M. W. (1982). Reciprocal effects of an inhibitory factor on catalytic activity and noncatalytic cGMP binding sites of rod phosphodiesterase. Proc. Natl. Acad. Sci. U. S. A. 79, 3702–3706. 144. Otto-Bruc, A., Antonny, B., Vuong, T. M., Chardin, P., Chabre, M. (1993). Interaction between the retinal cyclic GMP phosphodiesterase inhibitor and transducin. Kinetics and affinity studies. Biochemistry 32, 8636–8645. 145. Arshavsky, V. Y., Dumke, C. L., Zhu, Y., Artemyev, N. O., Skiba, N. P., Hamm, H. E., Bownds, M. D. (1994). Regulation of transducin GTPase activity in bovine rod outer segments. J. Biol. Chem. 269, 19882–19887. 146. Skiba, N. P., Artemyev, N. O., Hamm, H. E. (1995). The carboxyl terminus of the gammasubunit of rod cGMP phosphodiesterase contains distinct sites of interaction with the enzyme catalytic subunits and the α-subunit of transducin. J. Biol. Chem. 270, 13210– 13215. 147. Slepak, V. Z., Artemyev, N. O., Zhu, Y., Dumke, C. L., Sabacan, L., Sondek, J., Hamm, H. E., Bownds, M. D., Arshavsky, V. Y. (1995). An effector site that stimulates G-protein GTPase in photoreceptors. J. Biol. Chem. 270, 14319–14324. 148. Liu, Y., Arshavsky, V. Y., Ruoho, A. E. (1996). Interaction sites of the COOH-terminal region of the gamma subunit of cGMP phosphodiesterase with the GTP-bound α subunit of transducin. J. Biol. Chem. 271, 26900–26907. 149. Tsang, S. H., Burns, M. E., Calvert, P. D., Gouras, P., Baylor, D. A., Goff, S. P., Arshavsky, V. Y. (1998). Role for the target enzyme in deactivation of photoreceptor G protein in vivo. Science 282, 117–121. 150. Granovsky, A. E., Artemyev, N. O. (2001). A conformational switch in the inhibitory γ-subunit of PDE6 upon enzyme activation by transducin. Biochemistry 40, 13209–13215. 151. Whalen, M. M., Bitensky, M. W. (1989). Comparison of the phosphodiesterase inhibitory subunit interactions of frog and bovine rod outer segments. Biochem. J. 259, 13–19. 152. Rieke, F., Baylor, D. A. (1996). Molecular origin of continuous dark noise in rod photoreceptors. Biophys. J. 71, 2553–2572. 153. Wensel, T. G., Stryer, L. (1986). Reciprocal control of retinal rod cyclic GMP phosphodiesterase by its gamma subunit and transducin. Proteins 1, 90–99. 154. D’Amours, M. R., Cote, R. H. (1999). Regulation of photoreceptor phosphodiesterase catalysis by its noncatalytic cGMP binding sites. Biochem. J. 340, 863–869. 155. Yamazaki, A., Stein, P. J., Chernoff, N., Bitensky, M. W. (1983). Activation mechanism of rod outer segment cyclic GMP phosphodiesterase: release of inhibitor by the GTP/GTP binding protein. J. Biol. Chem. 258, 8188–8194. 156. Fersht, A. (1999). Structure and mechanism in protein science. Freeman, New York. 157. Dumke, C. L., Arshavsky, V. Y., Calvert, P. D., Bownds, M. D., Pugh, E. N., Jr. (1994). Rod outer segment structure influences the apparent kinetic parameters of cyclic GMP phosphodiesterase. J. Gen. Physiol. 103, 1071–1098. 158. Wensel, T. G., Stryer, L. (1990). Activation mechanism of retinal rod cyclic GMP phosphodiesterase probed by fluorescein-labeled inhibitory subunit. Biochemistry 29, 2155–2161. 159. Gillespie, P. G. (1990). Phosphodiesterases in visual transduction by rods and cones. In: Cyclic nucleotide phosphodiesterases: structure, regulation and drug action (Beavo, J., Houslay, M. D., eds.), pp. 163–184, Wiley, New York. 160. Whalen, M. M., Bitensky, M. W., Takemoto, D. J. (1990). The effect of the gamma-subunit of the cyclic GMP phosphodiesterase of bovine and frog (Rana catesbiana) retinal rod outer segments on the kinetic parameters of the enzyme. Biochem. J. 265, 655–658. 161. Melia, T. J., Malinski, J. A., He, F., Wensel, T. G. (2000). Enhancement of phototransduction protein interactions by lipid surfaces. J. Biol. Chem. 275, 3535–3542.

168

Cote

162. Bruckert, F., Catty, P., Deterre, P., Pfister, C. (1994). Activation of phosphodiesterase by transducin in bovine rod outer segments: Characteristics of the successive binding of two transducins. Biochemistry 33, 12625–12634. 163. Yamazaki, M., Li, N., Bondarenko, V. A., Yamazaki, R. K., Baehr, W., Yamazaki, A. (2002). Binding of cGMP to GAF domains in amphibian rod photoreceptor cGMP phosphodiesterase (PDE). J. Biol. Chem. 277, 40675–40686. 164. Phillips, W. J., Trukawinski, S., Cerione, R. A. (1989). An antibody-induced enhancement of the transducin-stimulated cyclic GMP phosphodiesterase activity. J. Biol. Chem. 264, 16679–16688. 165. Martins, T. J., Mumby, M. C., Beavo, J. A. (1982). Purification and characterization of a cyclic GMP-stimulated cyclic nucleotide phosphodiesterase from bovine tissues. J. Biol. Chem. 257, 1973–1979. 166. Yamamoto, T., Manganiello, V. C., Vaughan, M. (1983). Purification and characterization of cyclic GMP-stimulated cyclic nucleotide phosphodiesterase from calf liver. J. Biol. Chem. 258, 12526–12533. 167. Corbin, J. D., Turko, I. V., Beasley, A., Francis, S. H. (2000). Phosphorylation of phosphodiesterase-5 by cyclic nucleotide-dependent protein kinase alters its catalytic and allosteric cGMP-binding activities. Eur. J. Biochem. 267, 2760–2767. 168. Okada, D., Asakawa, S. (2002). Allosteric activation of cGMP-specific, cGMP-binding phosphodiesterase (PDE5) by cGMP. Biochem. 41, 9672–9679. 169. Rybalkin, S. D., Rybalkina, I. G., Shimizu-Albergine, M., Tang, X. B., Beavo, J. A. (2003). PDE5 is converted to an activated state upon cGMP binding to the GAF A domain. EMBO J. 22, 469–478. 170. Arshavsky, V. Y., Dumke, C. L., Bownds, M. D. (1992). Noncatalytic cGMP binding sites of amphibian rod cGMP phosphodiesterase control interaction with its inhibitory γsubunits. A putative regulatory mechanism of the rod photoresponse. J. Biol. Chem. 267, 24501–24507. 171. Yamazaki, A., Hayashi, F., Tatsumi, M., Bitensky, M. W., George, J. S. (1990). Interactions between the subunits of transducin and cyclic GMP phosphodiesterase in Rana catesbeiana rod photoreceptors. J. Biol. Chem. 265, 11539–11548. 172. Gillespie, P. G., Prusti, R. K., Apel, E. D., Beavo, J. A. (1989). A soluble form of bovine rod photoreceptor phosphodiesterase has a novel 15 kDa subunit. J. Biol. Chem. 264, 12187– 12193. 173. Korschen, H. G., Beyermann, M., Müller, F., Heck, M., Vantler, M., Koch, K. W., Kellner, R., Wolfrum, U., Bode, C., Hofmann, K. P., Kaupp, U. B. (1999). Interaction of glutamic-acidrich proteins with the cGMP signalling pathway in rod photoreceptors. Nature 400, 761–766. 174. Sugimoto, Y., Yatsunami, K., Tsujimoto, M., Khorana, H. G., Ichikawa, A. (1991). The amino acid sequence of a glutamic acid-rich protein from bovine retina as deduced from the cDNA sequence. Proc. Natl. Acad. Sci. U. S. A. 88, 3116–3119. 175. Colville, C. A., Molday, R. S. (1996). Primary structure and expression of the human βsubunit and related proteins of the rod photoreceptor cGMP-gated channel. J. Biol. Chem. 271, 32968–32974. 176. Batra-Safferling, R., Abarca, H. K., Korschen, H. G., Tziatzios, C., Stoldt, M., Budyak, I., Willbold, D., Schwalbe, H., Klein-Seetharaman, J., Kaupp, U. B. (2006). Glutamic acidrich proteins of rod photoreceptors are natively unfolded. J. Biol. Chem. 281, 1449–1460. 177. Pentia, D. C., Hosier, S., Cote, R. H. (2006). The glutamic acid-rich protein-2 (GARP2) is a high affinity rod photoreceptor phosphodiesterase (PDE6)-binding protein that modulates its catalytic properties. J. Biol. Chem. 281, 5500–5505.

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178. Poetsch, A., Molday, L. L., Molday, R. S. (2001). The cGMP-gated channel and related glutamic acid rich proteins interact with peripherin-2 at the rim region of rod photoreceptor disc membranes. J. Biol. Chem. 276, 48009–48016. 179. Florio, S. K., Prusti, R. K., Beavo, J. A. (1996). Solubilization of membrane-bound rod phosphodiesterase by the rod phosphodiesterase recombinant δ subunit. J. Biol. Chem. 271, 1–12. 180. Marzesco, A. M., Galli, T., Louvard, D., Zahraoui, A. (1998). The rod cGMP phosphodiesterase δ subunit dissociates the small GTPase Rab13 from membranes. J. Biol. Chem. 273, 22340–22345. 181. Zhang, H., Hosier, S., Terew, J. M., Zhang, K., Cote, R. H., Baehr, W. (2005). Assay and functional properties of PrBP(PDEδ), a prenyl binding protein interacting with multiple partners. Methods Enzymol. 403, 42–56. 182. Zhang, H. B., Liu, X. H., Zhang, K., Chen, C. K., Frederick, J. M., Prestwich, G. D., Baehr, W. (2004). Photoreceptor cGMP phosphodiesterase δ subunit (PDEδ) functions as a prenylbinding protein. J. Biol. Chem. 279, 407–413. 183. Zhang, H., Li, S., Doan, T., Rieke, F., Detwiler, P. B., Frederick, J. M., and Baehr, W. (2007). Deletion of PrBP/δ impedes transport of GRK1 and PDE6 catalytic subunits to photoreceptor outer segments. Proc. Natl. Acad. Sci. U. S. A. 104, 8857–8862. 184. Norton, A. W., Hosier, S., Terew, J. M., Li, N., Dhingra, A., Vardi, N., Baehr, W., Cote, R. H. (2005). Evaluation of the 17 kDa prenyl binding protein as a regulatory protein for phototransduction in retinal photoreceptors. J. Biol. Chem. 280, 1248–1256. 185. Hong, D. H., Pawlyk, B., Sokolov, M., Strissel, K. J., Yang, J., Tulloch, B., Wright, A. F., Arshavsky, V. Y., Li, T. (2003). RPGR isoforms in photoreceptor connecting cilia and the transitional zone of motile cilia. Invest. Ophthalmol. Vis. Sci. 44, 2413–2421. 186. Grayson, C., Bartolini, F., Chapple, J. P., Willison, K. R., Bhamidipati, A., Lewis, S. A., Luthert, P. J., Hardcastle, A. J., Cowan, N. J., Cheetham, M. E. (2002). Localization in the human retina of the X-linked retinitis pigmentosa protein RP2, its homologue cofactor C and the RP2 interacting protein Arl3. Hum. Mol. Genet. 11, 3065–3074. 187. Moritz, O. L., Tam, B. M., Hurd, L. L., Peränen, J., Deretic, D., Papermaster, D. S. (2001). Mutant rab8 impairs docking and fusion of rhodopsin-bearing post-Golgi membranes and causes cell death of transgenic Xenopus rods. Mol. Biol. Cell 12, 2341–2351. 188. Cook, T. A., Ghomashchi, F., Gelb, M. H., Florio, S. K., Beavo, J. A. (2000). Binding of the delta subunit to rod phosphodiesterase catalytic subunits requires methylated, prenylated C-termini of the catalytic subunits. Biochem. 39, 13516–13523. 189. Cook, T. A., Ghomashchi, F., Gelb, M. H., Florio, S. K., Beavo, J. A. (2001). The δ subunit of type 6 phosphodiesterase reduces light-induced cGMP hydrolysis in rod outer segments. J. Biol. Chem. 276, 5248–5255. 190. Coles, J. A., Yamane, S. (1975). Effects of adapting lights on the time course of the receptor potential of the anuran retinal rod. J. Physiol. (Lond.) 247, 180–207. 191. Calvert, P. D., Govardovskii, V. I., Arshavsky, V. Y., Makino, C. L. (2002). Two temporal phases of light adaptation in retinal rods. J. Gen. Physiol. 119, 129–146. 192. Krispel, C. M., Chen, C. K., Simon, M. I., Burns, M. E. (2003). Prolonged photoresponses and defective adaptation in rods of Gbeta5−/− mice. J. Neurosci. 23, 6965–6971. 193. Ebrey, T., Koutalos, Y. (2001). Vertebrate photoreceptors. Prog. Retin. Eye Res. 20, 49–94. 194. Zoraghi, R., Corbin, J. D., Francis, S. H. (2006). Phosphodiesterase-5 Gln817 is critical for cGMP, vardenafil, or sildenafil affinity: its orientation impacts cGMP but not cAMP affinity. J. Biol. Chem. 281, 5553–5558.

8 Rhodopsin Structure, Function, and Involvement in Retinitis Pigmentosa Scott Gleim and John Hwa CONTENTS Introduction Historical Perspective Rhodopsin as the Prototypical G Protein-Coupled Receptor Rhodopsin, Localization, and Signaling Dark State and Activation Structural Analysis Retinitis Pigmentosa Implications of Receptor Misfolding Nongenetic Contributions to RP Conclusion References

INTRODUCTION Rhodopsin is the dim-light sensitive photoreceptor, densely packed in the rod cells of the retina. Organisms from bacteria to humans have evolved highly specialized systems for the detection of light, driven by survival-based interests, ranging from energy capture to visual sensing. Phylogenic analysis suggests that photopic vision arose first as cone receptors, which diverged into four groups, with one of these groups further diverging to enable scotopic vision via rhodopsin [1]. Such an evolutionary scheme would suggest that highly sensitive dim-light photoreception developed through optimized specialization, that is, mutations surrounding the chromophore to refine photoactivation in terms of wavelength selectivity and, more importantly, sensitivity. The remarkable sensitivity of rhodopsin, activated by single photons, enables scotopic and peripheral vision. The 200-fs photoisomerization of rhodopsin remains among the fastest and most efficient biological photochemical reactions known. This capture of light energy and the corresponding visual response has mesmerized philosophers and scientists alike. From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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Fig. 1. A timeline for major events in rhodopsin research (1600 to present). Visual reception has a rich history. Investigations into vision are hinted at in writings from ancient Greece and in Egyptian images. A Systematic evaluation of visual system components required a philosophical revolution in scientific methodology. The nature of the eye (Kepler, 1604) and light optics (Newton, 1704) were among the earliest scientific investigations. Discovery of rhodopsin (Boll, 1876, and Kuehne, 1878) and the association of vitamin A deficiency with night blindness (Block, 1917, and Blegvad, 1924) paved the way for (B) the discovery of rhodopsin constituents (Wald, 1935). The following two decades of intense protein chemistry on rhodopsin led to a Nobel Prize for Wald (1967), stimulating new approaches to rhodopsin research and the identification of the opsin gene (Nathans, 1982–1985). These provided a vital tool for (C) detailed structural analyses, with spin-labeled interactions (Farrens, 1996, and Cai, 1997) and disulfide criticality (Karnik, 1988; Hwa, 1997; Hwa 2001) demonstrating important principles in activation and structure. Structural details of rhodopsin became visible by the seminal solution of the rhodopsin crystal structure (Palczewski, 2000). AFM atomic force microscopy, cGMP cyclic guanosine monophosphate, CSNB congenital stationary night blindness, ECM/EMC electron cryomicroscopy, EPR electron paramagnetic resonance, GPCR G protein-coupled receptor, ROS rod outer segment, RP retinitis pigmentosa.

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HISTORICAL PERSPECTIVE Systematic investigation into photoreception can be traced back to the years following the scientific revolution (Fig. 1A) when geographer, microscope enthusiast, and pioneer of microbiology Anton van Leeuwenhoek first observed retinal rod and cone cells in 1722, providing the first suggestion that light reception may occur somewhere other than at the lens. Thomas Young, famed for the double-slit experiment leading to the wave theory of light, subsequently proposed that color perception depends on three different color-sensitive nerve fibers, later defined by Hermann von Helmholtz to be blue, green, and red, a theory so advanced it was not proven until a century later. Heinrich Mueller suggested that retina rod and cone cells were involved in photoreception, further stimulating investigations into the retina. Franciscus Donders, around 1857, coined the term retinitis pigmentosa in a letter to Helmholz describing spicules of pigmentation he found throughout a patient’s degenerated retina. The discovery of rhodopsin is owed to the combined efforts of Franz Boll and Willy Kühne through an interesting series of exchanges reviewed elsewhere [2]. Bloch linked night blindness to malnutrition in 1917, from which Blegvad subsequently identified the deficient agent to be vitamin A. Involvement of vitamin A deficiency in night blindness provided supplemental evidence in the seminal identification of the active combination of vitamin A and opsin in 1935 by Wald (Fig. 1B), whose accomplishments have enabled incalculable benefits in vision research [3]. Over the past three decades, astounding progress has been made since the discovery of the opsin gene by Nathans [4]; these include the discovery of naturally occurring mutations leading to retinitis pigmentosa [5], deciphering the signaling pathway through transducin [6–12], determination of critical structural features such as the Schiff’s base counterion [13] and disulfide bond [14, 15], electron cryomicroscopy structure [16, 17], conformational movements required for activation [18, 19], and the crystal structure of rhodopsin [20] (Fig. 1C). Further details of activation are being intensely investigated at the molecular and biophysical levels [8, 21, 22]. Insights into the nature of misfolded rhodopsin [23–29] and dimeric packing of rhodopsin [30–34] have also been major achievements. This exponential rate of discovery will likely unfold many further details on the intriguing structure and function of the opsin protein and disease associations. RHODOPSIN AS THE PROTOTYPICAL G PROTEIN-COUPLED RECEPTOR Rhodopsin receives considerable research interest, particularly in structural and functional studies, owing in part to its reputation as the prototypical member of the seven-transmembrane-spanning, guanine nucleotide-binding protein (G protein)-coupled receptor superfamily, which accounts for approximately 5% of genes in the human genome. The G protein-coupled receptors (GPCRs) are arguably among the most medically important protein families as over 30% of available pharmaceuticals target proteins within this family [35]. Furthermore, rhodopsin remains the only crystallographic structure available to represent the GPCR superfamily. GPCRs are generally responsible for transmitting extracellular information into the cellular environment with ligand stimuli covering the range of biochemical diversity: large macromolecules, peptides, amino acids, nucleic acids, lipids, ions, and even, as with rhodopsin, a single photon of light.

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Much of what is known about rhodopsin, and thus GPCRs in general, is due in large part to studies performed on bovine rhodopsin. The bovine eye provides an ample source from which substantial amounts of this protein can be purified or studied directly. Sequencing of the bovine rhodopsin gene [4] gave an arrangement of five exons, subsequently identified (Fig. 1C) also to represent human rhodopsin gene configuration [36]. The 6.4-kb gene consists of a 96-bp 5' untranslated region; a 1,044-bp coding region; and a surprisingly long, approximately 1,400-bp 3' untranslated region and are divided into five exons by four introns that interrupt the coding region [4]. The human gene (Gene ID 6010) is located on chromosome 3 (3q21–q24). The resulting proteins are 93.4% homologous with completely conserved cytoplasmic loops. RHODOPSIN, LOCALIZATION, AND SIGNALING Expression of rhodopsin is required for normal cell morphology, as the rod outer segment (ROS) does not form in rhodopsin knockout mice (−/−) [37]. Interestingly, ROS formation takes on typical morphology in rhodopsin heterozygotes (+/−) [32], but with about 50–60% of typical ROS volume, decreased rhodopsin concentration, decreased 11-cis retinal concentration, and impaired light sensitivity [32, 37]. Rhodopsin is densely packed (most probably as dimers; [30]) into stacked disks within the ROS, constituting more than 90% of membrane protein in the lipid bilayer. Opsin is synthesized, folded, and transported through a nonmotile ciliary connection [38] between the cell body and the outer segment, where it functions as a G protein-coupled photon receptor. Disks are shed regularly, with the outermost ROS segments endocytosed by retinal pigment epithelia (RPE) and resulting vesicles trafficked to the RPE proteasomal compartment for opsin degradation. Isomerization of the retinal moiety, by light excitation, extends the twisted springlike carbon chain, forcing away nearby residues (Fig. 2). This conformational change exposes the hydrophobic binding site for a heterotrimeric G protein, transducin (Gt), to associate and become catalytically active, exchanging guanosine diphosphate (GDP) for guanosine triphosphate (GTP). In the case of rhodopsin, the GDP/GTP exchange dissociates Gt from opsin to bind phosphodiesterase, removing the inhibitory γ-subunits. This generates active cyclic guanosine monophosphate (cGMP)-phosphodiesterase, which in turn hydrolyses cGMP at a rate of 103 per second, rapidly closing cGMP-gated Na+ channels and hyperpolarizing the rod cell. Hyperpolarization stops neurotransmitter release, predominantly glutamate, to neighboring ganglia. A single GPCR activates multiple G proteins, which in turn activate multiple downstream signaling factors, resulting in a highly amplified signal of at least 10,000 hydrolyzed cGMP molecules per photon under dim-light conditions [39]. The signal is rapidly quenched through rhodopsin kinase (RK) phosphorylation of multiple serine and threonine residues along the C-terminal tail, allowing arrestin to bind and preventing further Gt interactions. These multiple phosphorylation sites appear to be critical to the remarkable reproducibility of the rhodopsin signal [40]. Arrestin binding promotes the hydrolysis and release of all-trans retinal, allowing for association of a new 11-cis retinal molecule, dependent on release of arrestin [41], thereby promoting dephosphorylation by protein phosphatase A (PPA) [42], regenerating a light-sensitive receptor.

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DARK STATE AND ACTIVATION Generally, opsins are large integral membrane proteins of approximately 360 amino acid residues, half comprising the GPCR characteristic seven-membrane-traversing regions (Fig. 3A). Despite countless similarities and conserved regions, rhodopsin is unique among this superfamily in a number of other ways. For instance, rhodopsin uniquely functions as a holoprotein, a working assembly of the precursor opsin apoprotein and a prosthetic inverse agonist, 11-cis retinal. This vitamin A derivative attaches covalently to lysine 296 through Schiff base formation, stabilizing opsin in a completely inactive conformation. This inverse agonist serves as the chromophore, finely tuning the absorption wavelength of the receptor in conjunction with surrounding residues. Such precision is required to prevent visual noise and allow for optimal visual sensitivity. In the absence of the 11-cis retinal, a significant degree of transducin coupling and thus signaling can occur. The spectrophotometric absorption profile of rhodopsin is defined by binding pocket interactions with the chromophore. This fortuitously allows tracking of structural changes by measuring the shift in the local absorption maxima (λmax) of the spectra. Dark-state rhodopsin maintains a characteristic absorption peak (λmax) at 498 nm, in which the bound ligand is maintained in a state reminiscent of a twisted spring. A photon of light energy strikes. On excitation, the positive charge, once localized to the Schiff base, redistributes along the π-electron system [43]. Charge transfer to an alternative counterion accompanies isomerization of the retinal molecule into all-trans retinal, sterically pushing apart transmembrane segments three (TM3) and six (TM6) [18]. Surrounding features of the holoprotein respond to the excitation in rapid succession, through a series of excited states, before final energy decay into a form relaxed enough to release all-trans retinal and activate the waiting effector, Gt. Bathorhodopsin (529 nm) develops equilibrium with a blue-shift intermediate (BSI) state (477 nm), which decays into a counterion transition state, lumirhodopsin (492 nm). Lumirhodopsin represents a transient state in which proton transfer from the E113 dark-state counterion [44] in TM3 across S186 and through an integral water molecule to protonate alternative counterion E181 [45] in the second intradiskal loop results in formation of meta I rhodopsin (478 nm). By the MI intermediate state, the ligand spring has untwisted, isomerized into all-trans retinylidene, still covalently attached to the opsin, and still incapable of activating transducin. Receptor activation, or conversion of meta I into meta II (MII or R*) (380 nm), requires deprotonation of the Schiff base, releasing the isomerized ligand.

Fig. 2. A Cartoon of the Rhodopsin Activation Cycle. A simplified rhodopsin activation scheme highlighting important events in the rhodopsin lifecycle. This schematic shows 11-cisretinal in (A) dark-state rhodopsin, with the protonated Schiff base stabilized by a counter-ion at Glu113. Photon energy catalyzes isomerization of the ligand to (B) all-trans-retinal, followed by counter-ion transfer across Ser186, during (C) lumi-rhodopsin, to Glu181, with a distancing of transmembrane helices 3 and 6 forming (D) active meta-II rhodopsin. Transducin (Gt) activation continues until (E) deactivating phosphorylation of the carboxy-tail, which promotes arrestin association and dissociation of all-trans-retinal. (F) Free opsin combines with new or re-formed 11-cis-retinal to reinitiate the cycling.

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Fig. 3. Structural representations of rhodopsin. (A) The primary sequence of rhodopsin, written as a schematic of the secondary structure, shows the organization of transmembrane helices. Autosomal dominant retinitis pigmentosa (RP) mutations are shown demonstrating the diverse range of affected residues and their locations. (B) A representation of rhodopsin, based on the 1LH9 2.6-Å crystal structure, highlights important structural features relevant to RP. Specifically, the amino (N-) and carboxy (C-) termini, the disulfide bridge, the retinal-binding site, and the highly conserved glutamate, arginine, tyrosine motif (ERY) activation-related sequence are associated with certain RP mutants.

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Accompanied rigid-body movements of TM3 and TM6 [46] expose a hydrophobic cleft between TM5 and TM6 [47]. Exposure of this hydrophobic site draws a nearby phenylalanine (F64) of the Gtγ, an orientation otherwise unfavored in an aqueous environment, resulting in amphipathic helix formation of the Gtγ-C-terminus, stabilized by activated rhodopsin, allowing allosteric regulation of nucleotide exchange [48].

STRUCTURAL ANALYSIS The current state of structural and functional knowledge regarding rhodopsin has been recently reviewed [49, 50]. Rhodopsin is ellipsoid in shape with dimensions of approximately 75 × 48 × 35 Å. The 348 amino acids of bovine rhodopsin are posttranslationally modified with N-terminus acetylation, N-terminus dual glycosylations (at N2 and N15), an intradiskal disulfide bond (C110–C187), dual palmitoylations at the C-terminus (C322 and C323), and multiple C-terminus light-activated phosphorylations. Electron Cryomicroscopy and Crystal Structure Most structural information on rhodopsin, and thereby on GPCRs, is based on an inactive, inverse-agonist-bound, dark state. This is because the most definitive structural data available are from x-ray crystallography. Electron cryomicroscopy leveraged twodimensional crystal formations to provide the earliest, albeit low-resolution, pictures of rhodopsin structure [16]. Originally solved to 2.8 Å [20], the crystal structure of rhodopsin demonstrated a number of important principles in rhodopsin function, the structure of GPCRs, and general aspects of large integral membrane proteins. Importantly, as mentioned, the helical arrangement was shown to be significantly different from, and more organized than, bacterial rhodopsin (another intensely studied seventransmembrane-spanning protein that serves as a proton pump from halophilic archaebacteria). Identification of proline-induced bending of transmembrane helices (e.g., highly conserved P267) showed significant distortion from an ideal helix, facilitating interhelical interactions and allowing for chromophore accommodation. Of particular interest was identification of a water-mediated interhelical interaction network centered around Asp83 on TM2, connecting this helix to TM3 and TM7 through interactions with Gly120 and Asn302, respectively. Also of mechanistic importance was the structural suggestion that β-ionone movement toward TM3 may result in helical displacement. Additional crystal analyses improved structural detail by adding missing residues (protein databank (http://rscb.org) structure identification numbers (pdf:1HZX), increasing resolution to 2.6 Å (1L9H) [51] (Fig. 3B), further improving resolution to 2.2 Å (1U19) [52], and refining earlier structures in different crystal space groups (1GZM). Increased resolving power provided confirmation of the structural importance of water molecules and their likely participation in spectral tuning and proton transfer [51]. Further details provided definition of the cytoplasmic region and chromophore, demonstrating a 6 s-cis conformation of the ionone ring and an unusual twisted and extended π-system with a delocalized charge–carboxylate interaction [52]. Also clarified was the hydrogen-bonding network connecting E113 to E181 through a required water molecule, a network later confirmed to transmit the counterion shift important in rhodopsin activation [45].

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Nuclear Magnetic Resonance Nuclear magnetic resonance (NMR) resolution of rhodopsin domain peptides have also been used to study the structure of rhodopsin [53]. Nuclear magnetic resonance has also extended crystallographic interpretations toward the elucidation of structural changes involved in opsin activation [54, 55]. Solid-state NMR, in particular, providing advantages for dealing with integral membrane proteins, is uniquely adept for following particular interactions, with studies utilizing isotopically labeled retinal providing direct chemical observation of modifications to ligand–protein interactions following light activation [55, 56]. The complete 1H and 13C assignments of the chromophore in the bound state showed interactions between 11-cis retinylidene’s H16/H17 and Phe 208, Phe 212, and H18 with Trp265 [57]. NMR provides considerable advantages in the context of activation as light-activated rhodopsin remains elusive to crystallization, and photoactivated intermediates, likely due to their transient nature, prove similarly difficult to crystallize [58]. The side chain of Glu122 and backbone of His211 were shown to be disrupted in meta II [55]. Cysteine Mutagenesis and Electron Paramagnetic Resonance Site-directed cysteine mutagenesis and sulfhydryl modification chemistry provide remarkable resources for structural studies facilitating spin labeling, disulfide construction [18, 59], and metal-binding site engineering [19]. Paramagnetic spin labeling adds unique topological information, measuring solvent accessibility of the modified residue, overall mobility of particular residues, and even demonstrating residue interactions. A series of spin-labeling studies revealed structural details of cytoplasmic loop 1 connecting helices 1 and 2 [60], loop 2 between helices 3 and 4 [12], loop 3 between helices 5 and 6 [61, 62], loop 4 leading from helix 7 to the palmitoylation site [63], and the C-terminal tail [64]. Together, these results map a range of light-initiated structural changes. Evaluating light-activated changes in mobility of cytoplasmic loop 3, the transducin-interacting domain connecting transmembrane helices 5 and 6, demonstrated a dramatic loss of mobility for residue V227, with a smaller decrease in mobility for K231. The loss of mobility corresponds to formation of tertiary contacts, whereas an increased mobility, as observed with V250, T251, M253, and Q244, indicates that tertiary contacts are lost in conversion to MII. This has considerable mechanistic importance as Q244 has been identified as a required residue for Gt activation [65]. As such, it can be seen that residues in cytoplasmic loop 3 are exposed to allow transducin interaction with nearby residues, forming new contacts to maintain structural integrity of the receptor. Other Approaches Breaking the protein into subsections for structural analysis of the components attempts to alleviate some of these concerns; however, less-direct biophysical measurements of intact protein have allowed structural inferences to fill gaps left by direct measurements. Fourier-transform infrared (FTIR) spectroscopy, for example, resolved proton movements involved in activation-induced counterion shift [44], and ultraviolet-visible (UV/Vis) spectrophotometry is routinely used to evaluate rhodopsin purity, structural stability, regeneration rate, and activation state [66]. Such techniques prove quite powerful

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at leveraging structural information provided by crystallographic data, particularly when combined with complementary tools such as site-directed mutagenesis and molecular modeling. In fact, molecular modeling has been pivotal in the study of rhodopsin, as it is a critical refining step in processing crystallographic and NMR data, and facilitates mutagenic approaches and biophysical data interpretation. Not surprisingly, as modeling techniques continue to mature, they become utilized as an experimental approach in their own right, with energy-induced decay of the protein structure revealing a core set of stabilizing interactions providing a folding scaffold for the overall rhodopsin structure [67]. Genetic manipulation techniques have proven useful in structural investigations of rhodopsin. From deletion of segments and chimeric recombination of protein to mutagenesis of individual residues and chemical modification of localized functional groups, each distinct application provides creative insight into structural features of this remarkable protein. Supporting a common GPCR activation principal is construction of a chimeric rhodopsin spliced with the cytoplasmic regions of the β-adrenergic receptor, resulting in a light-activated GPCR that elicits a β-adrenergic Gαs stimulation of cyclic adenosine monophosphate (cAMP) [68]. Investigations into rhodopsin structure and function parallel many major unanswered questions facing general protein biochemistry. Structural motions involved in translating binding of a ligand or allosteric modulator across the membrane bilayer to activate intracellular signals are of general interest, particularly if findings extrapolate to a wider range of GPCRs. More structurally accessible than most GPCRs, rhodopsin continues to provide a uniquely suited prototype for studying general GPCR features. One interesting feature to develop over recent years is the concept of GPCR dimerization, carrying uncertain potential impact [69, 70]. Rhodopsin has demonstrated potential to form dimers, as well as higher-order oligomers, in disk membranes [33], expression systems [71], liposomes [72] and in solubilizing detergents [34]. However, demonstration of native receptor dimerization, along with a functional requirement for dimerization, remains elusive. Perhaps the most compelling evidence in favor of rhodopsin dimer formation was demonstrated using atomic force microscopy [30]. A large battery of additional techniques, including electron microscopy, sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) separation, proteolysis, and cross-linking support the idea that rhodopsin is capable of dimer formation in both isolated disk membranes and when solubilized in detergent [33]. Site-directed mutagenesis, combined with fluorescence resonance energy transfer (FRET) and cysteine cross-linking, suggests a hydrophobic interaction between W175 in the second intradiskal loop, and Y206 in TM5 participates in rhodopsin dimer formation [71]. Molecular modeling has provided additional details for the putative interface [31]. Implications of native dimer formation for GPCRs range from transport considerations [73] to activation responses [74]. An interesting approach using solubilized rhodopsin in detergent micelles of increasing size accommodating mixtures of differing oligomeric sizes suggested that larger dimeric organizations might be structurally preferred, reflected by increased receptor stability [34]. Although these organizations may not be required for transducin activation, increasing levels of oligomerization corresponded to dramatically increased rates of Gt activation, without modifying MII decay, consistent with putatively improved Gt binding by dimeric rhodopsin [75].

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RETINITIS PIGMENTOSA Retinitis pigmentosa (RP) is a collection of inherited neurodegenerative disorders in which rod cell apoptosis spreads throughout the retina, resulting in progressive vision loss and the characteristic pigmented retinal appearance. RP genetics demonstrate extreme heterogeneity in severity, progression rate, and mode of inheritance, covering autosomal dominant, autosomal recessive, X-linked, and sporadic mutations. Over 30 different genes have been identified to produce RP, with additional loci also implicated. Of at least 15 autosomal dominant RP (adRP) inherited disease genes, some have obvious correlations to RP biochemistry, while others have uncertain or indeterminate relationship to the disease. By definition, all RP mutants result in photoreceptor degeneration. Degenerative effects trigger multiple cell-death pathways, including caspase-dependent apoptosis, complement activation, and autophagy [76], with phase profiles reflecting the initial cause of cell death, be it calcium overload, structural defects, or oxidative damage. Uniform retinal degeneration was demonstrated to be independent of cellular genotype in chimeric retinas [77], indicating that transcellular factors or interactions are responsible for the final global retinal degeneration. Rhodopsin mutations account for an estimated 40% of autosomal dominant gain-offunction mutations, with over 100 distinct mutations within the receptor (Fig. 3A). Initial association of RP with single-point mutations in rhodopsin stemmed from identification of the predominant rhodopsin RP mutant, P23H [5]. Clinical visual parameters of P23H RP suggest increased ROS shedding and impaired ROS renewal [78], findings consistent with the notion of misfolded opsin impairing the disk integrity. With an astounding number of different single-point mutations in rhodopsin leading to autosomal dominant RP [79], it becomes useful to categorize the resulting effects when evaluating disease mechanism and therapeutic approaches. Although the final pathology may appear to be similar, rhodopsin mutations are a heterogeneous group at the molecular level, with differing structural perturbations. Potential for pharmacological and molecular therapy may ultimately depend on the location of the mutation and amino acid change. Classification of RP mutants in terms of structural loci and mechanistic impact can be reduced to three critical structural regions [50]. Most severely, modifications in the cytoplasmic tail interfere with rhodopsin trafficking to the ROS. Mutants interfering with the normal disulfide bond formation, between C187 on the second intradiskal loop and C110 at the end of the first intradiskal loop, lead to receptor misfolding. Finally, mutations among the transmembrane- and chromaphore-binding regions can effect protein folding and receptor activation. Associating cytoplasmic, intradiskal, and transmembrane mutations with trafficking, misfolding, and activation defects provides a useful means to discuss major routes of rhodopsin-RP development, but also provides drastic overgeneralization of a highly heterogeneous problem. An extended classification scheme is based on inactivating mutations of GPCRs in general [79]: class I, defective biosynthesis; class II, defective trafficking; class III, defective ligand binding; class IV, defective activation; and class V, unknown defects. Defective biosynthesis predominantly occurs through premature termination, often through frameshift, but may also entail accelerated degradation. Defective surface trafficking, in this scheme, covers the majority of GPCR mutations, where normally produced receptors demonstrate intracellular retention. Receptors with ligand-binding defects,

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despite proper production and surface expression, are incompetent for ligand binding. Activation-defective receptors, it follows, demonstrate proper production, localization, and ligand binding, failing only to carry out the final activation step, resulting in reduced maximal response or sensitivity (manifest as an increased half maximal response, EC50). The final category of unknown defects refers to disease-associated mutants with no apparent mechanistic deficiency, functioning normally in model systems and suggesting a case for which, despite association, the mutant may not be the cause of disease. While categorizing mutation effects may facilitate discussion of general mechanisms, the differences in location and physicochemical properties of replacement residues likely result in a broad range of subtle changes among each major class. Mutation of the arginine in the conserved [E/D]RY sequence associated with G protein signaling results in impaired activation in the melanocortin MC1R receptor (R142H) through G protein decoupling. The corresponding mutation in the vasopressin V2R receptor, R147H, also causes constitutive internalization and desensitization [80], resulting in nephrogenic diabetes insipidus. Similar RP mutants have been found, R135L and R135W, that are unable to activate transducin despite normal folding and ligand binding [81], a phenotype that could fall into the class IV activation-impaired mutant group. However, further research into these mutants has identified the defect to be caused by constitutive phosphorylation and arrestin binding, leading to constitutive internalization [82]. One could also argue that this may fall into the category of defective trafficking. However, the cellular outcome of this defect remains distinctly different from that of traditional mistrafficking. Transmembrane RP Rhodopsin Mutants The transmembrane region of rhodopsin, consisting of half of the protein, is of obvious structural importance. This region also forms the retinal-binding pocket and coordinates G protein activation through conformational movements of the transmembrane (TM) domains. As such, rhodopsin mutations within the transmembrane region can be further subdivided by those in proximity of the retinal β-ionone ring or carbon chain and those found in remaining regions of the transmembrane domains (Fig. 4). Evaluation of RP mutants across the transmembrane domains (G51A,V in TM1, G89D in TM2, L125R,A,F in TM3, A164V in TM4, H211P in TM5, P267L,R in TM6, and T297R in TM7) demonstrated that mutations in each TM segment can lead to abnormal bleaching and MII photointermediates [25], indicators of protein misfolding. These mutations appear to result in nonnative packing of the transmembrane helices, which relay misfolding to the intradiskal domain, where they may cause abnormal disulfide formation. The L125 residue is in TM3, within the ligand-binding pocket, close to the retinal β-ionone ring. A comprehensive list of mutations at this site (G, N, I, H, P, T, D, E, Y, and W) decreased 11-cis retinal binding, causing a red-shift of λmax, increased solvent exposure, and decreased thermostability [24, 83]. These findings support the importance of ligand binding in maintaining the structural integrity of rhodopsin, highlighting an interaction between L125 and the β-ionone ring as critical in maintaining the structure of the chromaphore-binding pocket. Further use of mutagenesis in evaluating the structural role of L125 demonstrated the role of this residue in maintaining additional interhelical interactions [29]. Rescue of the L125R RP mutant through compensatory mutations of W126E,D or E122L eliminated steric hindrance caused by L125R, restoring

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Fig. 4. (continued)

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Fig. 4. Regional clusters of autosomal dominant retinitis pigmentosa (adRP) rhodopsin mutations. Organization of adRP mutations according to structural features demonstrates clusters in relation to (A) the retinal-binding site, (B) the transmembrane scaffold, (C) the cytoplasmic region, and (D) the intradiskal region.

the TM3–TM5 interaction formed by the salt bridge between E122 and H211, by reconstructing the hydrogen bond between W126 and E122. Similarly, RP mutant A164 in TM4 interfered with residues L119 and I123 in TM3, disrupting the same salt bridge. Congenital stationary night blindness (CSNB; nyctalopia) is a condition characterized by inability to see in conditions of low illumination. The most common cause of nyctalopia

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is rod photoreceptor degeneration from RP, often caused by rhodopsin mutations. Non-RP mutations generating constitutive receptor activation cause CSNB. Rhodopsin is constrained in an inactive conformation through the salt-bridge interaction between lysine-296 and glutamate-113 [84]; mutation of either component can generate constitutive activity, causing CSNB. Interestingly, one of the earliest symptoms of RP is night blindness [85], demonstrating a fascinating relationship between rhodopsin folding and activity. RP mutation K296E, however, has been shown to cause photoreceptor degeneration through a process independent of constitutive activity [86]. Proximity of glycine-90 to this counterion permits G90D interference of the K296-E113 salt bridge by substituting for E113 in salt-bridge formation, leading to constitutive activation and CSNB [87]. The T94I autosomal dominant CSNB mutation is similarly near the G90 region and also results in constitutive transducin activation, most likely through hydrophobic interference or steric hindrance [88]. Cytoplasmic RP Rhodopsin Mutants The cytoplasmic region of rhodopsin extends from the surface of disks into the cellular milieu, where transducin (Gt) binding, and hence signaling, occurs. This intracellular region consists of the first cytoplasmic loop connecting helices 1 and 2, the second cytoplasmic loop connecting helices 3 and 4, the third cytoplasmic loop, and the C-terminal tail. The C-terminus is anchored to the membrane surface by palmitoylation of residues Cys322 and Cys323, creating what is putatively referred to as the fourth cytoplasmic loop [89, 90]. The proximal portion of this loop forms a part of the binding site for the C-terminal section of the transducin α-subunit [91]. RP mutations within the cytoplasmic region of rhodopsin are found throughout the region, with the exception of the third cytoplasmic loop (Fig. 4C). The C-terminus directly interacts with a cargo-binding subunit of dynein, providing transport of rhodopsin to the outer segment [92], an interaction abolished by severe RP mutants P347L, P347S, V345M, and Q344ter. C-terminal tail also contains multiple phosphorylation sites, phosphorylated by RK following transducin activation. Phosphorylation of the C-terminal tail regulates interaction of rhodopsin with arrestin, deactivating the receptor. These multiple phosphorylation sites result in uniquely consistent control of the amplitude and duration of the activated rhodopsin signal [40]. Intradiskal RP Rhodopsin Mutants Structural investigations of the RP mutants and their consequences on the intradiskal region were best highlighted in a series of investigations by Khorana et al. [23, 25, 26, 93–95]. Investigations by this group established a number of techniques critical for furthering rhodopsin research, while demonstrating important general principles of rhodopsin structure and the misfolding properties of numerous RP mutants. Together, their findings suggest that packing of the transmembrane helices, binding of the chromaphore, and intradiskal structural integrity are physically coupled properties critical for proper folding of rhodopsin. Intradiskal RP mutations are numerous and diverse in their impact on misfolding (Fig. 4D). Deletion studies of the intradiskal region provided suggestions for the structural importance of this region [96]. N-terminal deletions resulted in partial chromophore

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regeneration, whereas deletions in the first and second intradiskal loops caused regenerationincompetent misfolding, with segments 171–182 and 189–192 being structurally essential. Single-point mutations utilized to investigate the relative contribution of residues in the deleted segments along the second intradiskal loop showed that most individual changes resembled wild-type regeneration. Deletion of 189–190 resulted in dramatic retention of the mutant protein in the endoplasmic reticulum. This region is now known to contribute to the formation of a structurally critical salt bridge between D190 and R177 [97], linking the ends of the second intradiskal loop. Although interaction of this ion pair has no effect on solvent exposure or signaling, this interaction appears conserved as R or K and E or D pairings in most GPCRs and is critical for stabilization of the dark state of rhodopsin. The stabilizing role of this salt bridge is highlighted by its proximity to the disulfide bond between C110 and C187 in the intradiskal region. Formation of this disulfide bridge is critical to receptor structure and is highly conserved throughout the GPCR superfamily. RP mutants directly modifying one of these cysteines, C110F and C110Y or C187Y, cause abnormal disulfide formation of C185:C187 or C110:C185, respectively [26]. Surprisingly, additional RP mutants (G89D, L125R, A164V, and H211P) influence this structural bridge, through disruption of normal helical packing, promoting formation of the abnormal C185:C187 disulfide bridge [15]. IMPLICATIONS OF RECEPTOR MISFOLDING The dominant impact of misfolded rhodopsin is demonstrated through coexpression of wild-type rhodopsin and a misfolded mutant, for which intracellular retention of the mutant results in a corresponding retention of wild-type receptor, decreasing surface expression and signaling [79]. Numerous structural elements are ultimately responsible for maintenance of the rhodopsin structure, including membrane lipid interactions [98, 99], salt bridges, and both hydrophobic and polar contacts within the transmembrane regions. However, no single interaction has demonstrated as much significance in stabilization of the overall receptor as the conserved disulfide bond located at the intradiskal–transmembrane interface. Evaluating the rate of vision decline among 140 RP patients from 1975 to 2000 showed C-terminal rhodopsin mutations to decline most rapidly [100]. Considering the various classifications of rhodopsin mutations eliciting RP, it seems intuitive that significant variability in disease severity and rate of progression would arise between the various classes. Indeed, adRP mutations in rhodopsin show astounding variability within a given mutation class [85], even when considering only a single-point mutation, such as P23H. The progression model included age, gender, baseline function, and affected region but could only account for 20–34% of the variation. Predominant theories behind individual variability in disease progression focus on complementary genetic and nongenetic contributions. Complementary genetic contributions may include any polygenic interaction affecting retina function, including rhodopsin folding, trafficking, degradation, ion homeostasis, RPE integrity, ROS structure, and vitamin A processing. Detailed analyses involving polygenic interactions in RP are becoming more accessible with advances in techniques to identify single-nucleotide polymorphisms and perform haplotype analysis, with findings anticipated to have considerable impact on disease severity and progression.

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Acknowledging differences between the various forms of RP may lead to interesting implications in the prospect of treatments. Misfolding and aberrant disulfide formation, for example, may be alleviated through traditional pharmacological intervention with retinoid-mimetic folding chaperones. Transgenic mice expressing the misfolding T17M mutation and receiving dietary supplementation with high-dose vitamin A showed significant reduction in degeneration symptoms [101]. No such effect was observed in transgenic mice expressing the trafficking-impaired P347S variant, suggesting that such a therapeutic approach would indeed be specific to misfolding. In vitro, similar receptor rescue improves purification yield of severely misfolded RP mutant A164V using increased concentrations of inverse agonist [66]. The most predominant rhodopsin RP mutation, P23H, is also a misfolding mutation, resulting in intracellular accumulation [102]. Use of retinal-based structural chaperones 9-cis retinal, 11-cis retinal, and 11-cis 7-ring retinal [103] improved the ability of the photoreceptor to reach the plasma membrane [104], although not necessarily resulting in functional protein production [105]. Demethylation at C1 or C5 of the retinal ring produces partial agonist activity, shifting conformational equilibrium from MII to inactive MI by interfering with E134mediated proton transfer [106]. Thus, pharmacotherapy for rhodopsin RP mutations may require a degree of individualization based on specific structural defects imparted by the mutations. Simple in vitro assays may be of potential use in predicting in vivo responsiveness. NONGENETIC CONTRIBUTIONS TO RP Nongenetic contributions are also being elucidated. Suggested factors affecting the clinical course of RP focus primarily on diet, general health, and light exposure. The observation that light exposure exacerbates retinal degeneration in particular RP subtypes provides an interesting and complicating factor in understanding disease progression. One explanation for this phenomenon relates to cellular damage incurred by ultraviolet radiation, potentially damaging the rod cell, the RPE, or other surrounding supportive cell types. An alternative, and potentially additive, hypothesis suggests that ligand, released on photoactivation, becomes unable to stabilize mutant opsin, allowing the receptor to collapse and encouraging the misfolded-degeneration process. The theory of exacerbated misfolding/instability due to loss of ligand is indirectly supported by the only common treatment available for RP patients, vitamin A supplementation. As a required precursor to 11-cis retinal formation, which acts as a structural stabilizing agent for rhodopsin, treatment with vitamin A provides support for mutations affecting protein folding. However, effectiveness is, again, highly variable. One likely explanation for the variability of effectiveness with vitamin A supplementation in RP patients is the diversity of causal defects, as described with rhodopsin mutation classifications, which would be expected to produce various responses. Precisely such a difference in effectiveness was demonstrated through comparison of vitamin A effectiveness in transgenic mice expressing either a trafficking-impaired mutant (P347S) or a foldingimpaired mutant (T17M) [101]. As anticipated, based on the stabilizing effect of ligand binding, vitamin A improved histologic morphology and decreased the rate of decline for the misfolding mutant but not for the trafficking-impaired mutant.

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Circumstantial evidence points to another supplement with potential impact on disease progression. Association of zinc deficiency with a number of clinical manifestations of RP, impaired dark adaptation [107], decreased rhodopsin regeneration [108], and degeneration of ROS [109], has suggested possible involvement of the essential trace metal in both native functional and pathological roles. Treatment of bullfrog eyes with low-level zinc resulted in elevated dark-adapted electroretinogram (ERG) thresholds, increased peak ERG amplitudes, and accelerated rhodopsin regeneration [110]. As a critical component of the retina, zinc concentrations in ROS extracts suggest that it may be fortified in ROS disks [111], and radionuclide 65Zn has shown direct binding to purified rhodopsin [112]. Recent evidence also supports direct association of zinc at a high-affinity coordination site near H211 and E122 [113] as well as concentration-dependent alteration of rhodopsin thermostability [114]. Crystallographic data also suggest the presence of such a site [20], although this is not reflected in some later structures as the resolved metal ions were manually replaced with waters during processing [52]. Although solid-state NMR data suggest a lack of direct binding between H211 and zinc, the observed chemical shifts are consistent with the presence of Zn2+ within that region [115]. Direct interactions between divalent cations and GPCRs are not unprecedented as a diverse list of receptors, including β2-adrenergic [116, 117], dopaminergic [118], melanocortin MC1 and MC4 [119, 120], and olfactory receptors [121], have demonstrated direct and specific interactions with zinc. Evaluation of zinc and other trace metals in the context of RP has remained suggestive, although inconclusive [122–124], likely due to a focus on serum levels and a lack of stratification based on different RP mechanisms. CONCLUSION The development of powerful biochemical and biophysical techniques to study rhodopsin has allowed for considerable advances and understanding of this fascinating photoreceptor protein. The heterogeneous RP mutations have provided additional insights into rhodopsin structure/function. In vitro biochemical and biophysical assays will provide valuable information on potential therapy at the molecular level. Improvements with vitamin A treatment could be predicted through 11-cis retinal regeneration assays, and the effects of trace metals on folding and stability could also be evaluated. Comprehensive biochemical correlation to clinical disease is a necessary direction for future studies. One can envision as new mutations are discovered that such simple characterization of the mutants will determine appropriate therapy and may predict disease progress and prognosis. REFERENCES 1. Okano T, Kojima D, Fukada Y, Shichida Y, Yoshizawa T. Primary structures of chicken cone visual pigments: vertebrate rhodopsins have evolved out of cone visual pigments. Proc Natl Acad Sci U S A 1992;89:5932–5936. 2. Marmor MF, Martin LJ. 100 years of the visual cycle. Survey Ophthalmol 1978;22:279–285. 3. Wald G. Molecular basis of visual excitation. Science 1968;162:230–239. 4. Nathans J, Hogness D. Isolation, sequence analysis, and intron-exon arrangement of the gene encoding bovine rhodopsin. Cell 1983;34:807–814.

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5. Dryja TP, McGee TL, Reichel E, et al. A point mutation of the rhodopsin gene in one form of retinitis pigmentosa. Nature 1990;343:364–366. 6. Fung BK, Hurley JB, Stryer L. Flow of information in the light-triggered cyclic nucleotide cascade of vision. Proc Natl Acad Sci U S A 1981;78:152–156. 7. Bennett N, Dupont Y. The G-protein of retinal rod outer segments (transducin). Mechanism of interaction with rhodopsin and nucleotides. J Biol Chem 1985;260:4156–4168. 8. Ridge KD, Marino JP, Ngo T, Ramon E, Brabazon DM, Abdulaev NG. NMR analysis of rhodopsin-transducin interactions. Vision Res 2006;45:4482–4492. 9. Herrmann R, Heck M, Henklein P, Hofmann KP, Ernst OP. Signal transfer from GPCRs to G Proteins: role of the G{alpha} N-terminal region in rhodopsin-transducin coupling. J Biol Chem 2006;281:30234–30241. 10. Wilden U, Wust E, Weyand I, Kuhn H. Rapid affinity purification of retinal arrestin (48 kDa protein) via its light-dependent binding to phosphorylated rhodopsin. FEBS Lett 1986;207:292–295. 11. Konig B, Arendt A, McDowell JH, Kahlert M, Hargrave PA, Hofmann KP. Three cytoplasmic loops of rhodopsin interact with transducin. Proc Natl Acad Sci U S A 1989;86:6878–6882. 12. Farahbakhsh ZT, Ridge KD, Khorana HG, Hubbell WL. Mapping light-dependent structural changes in the cytoplasmic loop connecting helixes c and d in rhodopsin: a site-directed spin labeling study. Biochemistry 1995;34:8812–8819. 13. Sakmar TP, Franke RR, Khorana HG. Glutamic acid-113 serves as the retinylidene schiff base counterion in bovine rhodopsin. Proc Natl Acad Sci U S A 1989;86:8309–8313. 14. Karnik SS, Sakmar TP, Chen HB, Khorana HG. Cysteine residues 110 and 187 are essential for the formation of correct structure in bovine rhodopsin. Proc Natl Acad Sci U S A 1988;85:8459–8463. 15. Hwa J, Klein-Seetharaman J, Khorana HG. Structure and function in rhodopsin: mass spectrometric identification of the abnormal intradiscal disulfide bond in misfolded retinitis pigmentosa mutants. Proc Natl Acad Sci U S A 2001;98:4872–4876. 16. Unger VM, Schertler GF. Low resolution structure of bovine rhodopsin determined by electron cryo-microscopy. Biophys J 1995;68:1776–1786. 17. Unger VM, Hargrave PA, Baldwin JM, Schertler GFX. Arrangement of rhodopsin transmembrane [alpha]-helices. Nature 1997;389:203–206. 18. Farrens DL, Altenbach C, Yang K, Hubbell WL, Khorana HG. Requirement of rigid-body motion of transmembrane helices for light activation of rhodopsin. Science 1996;274: 768–770. 19. Sheikh SP, Zvyaga TA, Lichtarge O, Sakmar TP, Bourne HR. Rhodopsin activation blocked by metal-ion-binding sites linking transmembrane helices C and F. Nature 1996;383: 347–350. 20. Palczewski K, Kumasaka T, Hori T, et al. Crystal Structure of rhodopsin: a g protein-coupled receptor. Science 2000;289:739–745. 21. Downs MA, Arimoto R, Marshall GR, Kisselev OG. G-protein alpha and beta-gamma subunits interact with conformationally distinct signaling states of rhodopsin. Vision Res 2006;46:442–448. 22. Vogel R, Siebert F, Yan ECY, Sakmar TP, Hirshfeld A, Sheves M. Modulating rhodopsin receptor activation by altering the pka of the retinal Schiff base. J Am Chem Soc 2006;128:10503–10512. 23. Liu X, Garriga P, Khorana HG. Structure and function in rhodopsin: correct folding and misfolding in two point mutants in the intradiscal domain of rhodopsin identified in retinitis pigmentosa. Proc Natl Acad Sci U S A 1996;93:4554–4559.

Rhodopsin and Retinitis Pigmentosa

191

24. Garriga P, Liu X, Khorana HG. Structure and function in rhodopsin: correct folding and misfolding in point mutants at and in proximity to the site of the retinitis pigmentosa mutation Leu-125→Arg in the transmembrane helix C. Proc Natl Acad Sci U S A 1996;93: 4560–4564. 25. Hwa J, Garriga P, Liu X, Khorana HG. Structure and function in rhodopsin: packing of the helices in the transmembrane domain and folding to a tertiary structure in the intradiscal domain are coupled. Proc Natl Acad Sci U S A 1997;94:10571–10576. 26. Hwa J, Reeves PJ, Klein-Seetharaman J, Davidson F, Khorana HG. Structure and function in rhodopsin: further elucidation of the role of the intradiscal cysteines, Cys-110, -185, and -187, in rhodopsin folding and function. Proc Natl Acad Sci U S A 1999;96:1932–1935. 27. Hwa J, Klein-Seetharaman J, Khorana HG. Structure and function in rhodopsin: mass spectrometric identification of the abnormal intradiscal disulfide bond in misfolded retinitis pigmentosa mutants. Proc Natl Acad Sci U S A 2001;98:4872–4876. 28. Ramon E, del Valle LJ, Garriga P. Unusual thermal and conformational properties of the rhodopsin congenital night blindness mutant Thr-94→Ile. J Biol Chem 2003;278:6427–6432. 29. Stojanovic A, Hwang I, Khorana HG, Hwa J. Retinitis pigmentosa rhodopsin mutations L125R and A164V perturb critical interhelical interactions: new insights through compensatory mutations and crystal structure analysis. J Biol Chem 2003;278:39020–39028. 30. Fotiadis D, Liang Y, Filipek S, Saperstein DA, Engel A, Palczewski K. Atomic-force microscopy: rhodopsin dimers in native disc membranes. Nature 2003;421:127–128. 31. Fotiadis D, Liang Y, Filipek S, Saperstein DA, Engel A, Palczewski K. The G proteincoupled receptor rhodopsin in the native membrane. FEBS Lett Struct Dyn Funct Proteins Biol Membr 2004;564:281–288. 32. Liang Y, Fotiadis D, Maeda T, et al. Rhodopsin signaling and organization in heterozygote rhodopsin knockout mice. J Biol Chem 2004;279:48189–48196. 33. Suda K, Filipek S, Palczewski K, Engel A, Fotiadis D. The supramolecular structure of the GPCR rhodopsin in solution and native disc membranes. Mol Membr Biol 2004;21: 435–446. 34. Jastrzebska B, Fotiadis D, Jang GF, Stenkamp RE, Engel A, Palczewski K. Functional and structural characterization of rhodopsin oligomers. J Biol Chem 2006;281:11917–11922. 35. Schlyer S, Horuk R. I want a new drug: G-protein-coupled receptors in drug development. Drug Discovery Today 2006;11:481–493. 36. Nathans J, Hogness DS. Isolation and nucleotide sequence of the gene encoding human rhodopsin. Proc Natl Acad Sci U S A 1984;81:4851–4855. 37. Lem J, Krasnoperova NV, Calvert PD, et al. Morphological, physiological, and biochemical changes in rhodopsin knockout mice. Proc Natl Acad Sci U S A 1999;96:736–741. 38. Eley L, Yates LM, Goodship JA. Cilia and disease. Current Opin Genet Dev 2005;15:308– 314. 39. Ames A 3d, Walseth T, Heyman R, Barad M, Graeff R, Goldberg N. Light-induced increases in cGMP metabolic flux correspond with electrical responses of photoreceptors. J Biol Chem 1986;261:13034–13042. 40. Doan T, Mendez A, Detwiler PB, Chen J, Rieke F. Multiple phosphorylation sites confer reproducibility of the rod’s single-photon responses. Science 2006;313:530–533. 41. Hofmann K, Pulvermuller A, Buczylko J, Van Hooser P, Palczewski K. The role of arrestin and retinoids in the regeneration pathway of rhodopsin. J Biol Chem 1992;267:15701– 15706. 42. Palczewski K, Hargrave PA, McDowell JH, Ingebritsen TS. The catalytic subunit of phosphatase 2A dephosphorylates phosphoopsin. Biochemistry 1989;28:415–419.

192

Gleim and Hwa

43. Mathies R, Stryer L. Retinal has a highly dipolar vertically excited singlet state: implications for vision. Proc Natl Acad Sci U S A 1976;73:2169–2173. 44. Jager F, Fahmy K, Sakmar TP, Siebert F. Identification of glutamic acid 113 as the Schiff base proton acceptor in the metarhodopsin II photointermediate of rhodopsin. Biochemistry 1994;33:10878–10882. 45. Yan ECY, Kazmi MA, Ganim Z, et al. Retinal counterion switch in the photoactivation of the G protein-coupled receptor rhodopsin. Proc Natl Acad Sci U S A 2003;100:9262–9267. 46. Farrens DL, Altenbach C, Yang K, Hubbell WL, Khorana HG. Requirement of rigid-body motion of transmembrane helices for light activation of rhodopsin. Science 1996;274:768–770. 47. Janz JM, Farrens DL. Rhodopsin activation exposes a key hydrophobic binding site for the transducin {alpha}-subunit C terminus. J Biol Chem 2004;279:29767–29773. 48. Kisselev OG, Downs MA. Rhodopsin controls a conformational switch on the transducin [gamma] subunit. Structure 2003;11:367–373. 49. Palczewski K. G Protein&#×2013;coupled receptor rhodopsin. Annu Rev Biochem 2006;75:743–767. 50. Stojanovic A, Hwa J. Rhodopsin and retinitis pigmentosa: shedding light on structure and function. Recept Channels 2002;8:33–50. 51. Okada T, Fujiyoshi Y, Silow M, Navarro J, Landau EM, Shichida Y. Functional role of internal water molecules in rhodopsin revealed by x-ray crystallography. Proc Natl Acad Sci U S A 2002;99:5982–5987. 52. Okada T, Sugihara M, Bondar A-N, Elstner M, Entel P, Buss V. The retinal conformation and its environment in rhodopsin in light of a new 2.2 A crystal structure. J Mol Biol 2004;342:571–583. 53. Yeagle PL, Albert AD. Structure of the G-protein-coupled receptor, rhodopsin: a domain approach. Biochem Soc Trans 1998;26:520–531. 54. Klein-Seetharaman J, Getmanova EV, Loewen MC, Reeves PJ, Khorana HG. NMR spectroscopy in studies of light-induced structural changes in mammalian rhodopsin: applicability of solution (19)F NMR. Proc Natl Acad Sci U S A 1999;96:13744–13749. 55. Patel AB, Crocker E, Reeves PJ, et al. Changes in interhelical hydrogen bonding upon rhodopsin activation. J Mol Biol 2005;347:803–812. 56. Smith SO, Palings I, Copie V, et al. Low-temperature solid-state 13C NMR studies of the retinal chromophore in rhodopsin. Biochemistry 1987;26:1606–1611. 57. Creemers AF, Kiihne S, Bovee-Geurts PH, DeGrip WJ, Lugtenburg J, de Groot HJ. (1)H and (13)C MAS NMR evidence for pronounced ligand-protein interactions involving the ionone ring of the retinylidene chromophore in rhodopsin. Proc Natl Acad Sci U S A 2002;99:9101–9106. 58. Spooner PJ, Sharples JM, Goodall SC, et al. Conformational similarities in the beta-ionone ring region of the rhodopsin chromophore in its ground state and after photoactivation to the metarhodopsin-I intermediate. Biochemistry 2003;42:13371–13378. 59. Cai K, Klein-Seetharaman J, Altenbach C, Hubbell WL, Khorana HG. Probing the dark state tertiary structure in the cytoplasmic domain of rhodopsin: proximities between amino acids deduced from spontaneous disulfide bond formation between cysteine pairs engineered in cytoplasmic loops 1, 3, and 4. Biochemistry 2001;40:12479–12485. 60. Altenbach C, Klein-Seetharaman J, Hwa J, Khorana HG, Hubbell WL. Structural features and light-dependent changes in the sequence 59–75 connecting helices I and II in rhodopsin: a site-directed spin-labeling study. Biochemistry 1999;38:7945–7949. 61. Altenbach C, Yang K, Farrens DL, Farahbakhsh ZT, Khorana HG, Hubbell WL. Structural features and light-dependent changes in the cytoplasmic interhelical E-F loop region of rhodopsin: a site-directed spin-labeling study. Biochemistry 1996;35:12470–12478.

Rhodopsin and Retinitis Pigmentosa

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62. Yang K, Farrens DL, Hubbell WL, Khorana HG. Structure and function in rhodopsin. Single cysteine substitution mutants in the cytoplasmic interhelical E-F loop region show positionspecific effects in transducin activation. Biochemistry 1996;35:12464–12469. 63. Altenbach C, Cai K, Khorana HG, Hubbell WL. Structural features and light-dependent changes in the sequence 306–322 extending from helix VII to the palmitoylation sites in rhodopsin: a site-directed spin-labeling study. Biochemistry 1999;38:7931–7937. 64. Langen R, Cai K, Altenbach C, Khorana HG, Hubbell WL. Structural features of the Cterminal domain of bovine rhodopsin: a site-directed spin-labeling study. Biochemistry 1999;38:7918–7924. 65. Yang K, Farrens DL, Hubbell WL, Khorana HG. Structure and function in rhodopsin. Single cysteine substitution mutants in the cytoplasmic interhelical E-F loop region show positionspecific effects in transducin activation. Biochemistry 1996;35:12464–12469. 66. Reeves PJ, Hwa J, Khorana HG. Structure and function in rhodopsin: kinetic studies of retinal binding to purified opsin mutants in defined phospholipid-detergent mixtures serve as probes of the retinal binding pocket. Proc Natl Acad Sci U S A 1999;96:1927–1931. 67. Rader AJ, Anderson G, Isin B, Khorana HG, Bahar I, Klein-Seetharaman J. Identification of core amino acids stabilizing rhodopsin. Proc Natl Acad Sci U S A 2004;101:7246–7251. 68. Kim J-M, Hwa J, Garriga P, Reeves PJ, RajBhandary UL, Khorana HG. Light-driven activation of beta2-adrenergic receptor signaling by a chimeric rhodopsin containing the beta2adrenergic receptor cytoplasmic loops. Biochemistry 2005;44:2284–2292. 69. Fotiadis D, Jastrzebska B, Philippsen A, Muller DJ, Palczewski K, Engel A. Structure of the rhodopsin dimer: a working model for G-protein-coupled receptors. Current Opin Struct Biol 2006;16:252–259. 70. Angers S, Salahpour A, Bouvier M. Dimerization: an emerging concept for G protein-coupled receptor ontogeny and function. Annu Rev Pharmacol Toxicol 2002;42:409–435. 71. Kota P, Reeves PJ, Rajbhandary UL, Khorana HG. Opsin is present as dimers in COS1 cells: identification of amino acids at the dimeric interface. Proc Natl Acad Sci U S A 2006;103:3054–3059. 72. Mansoor SE, Palczewski K, Farrens DL. Rhodopsin self-associates in asolectin liposomes. Proc Natl Acad Sci U S A 2006;103:3060–3065. 73. Jordan BA, Trapaidze N, Gomes I, Nivarthi R, Devi LA. Oligomerization of opioid receptors with beta 2-adrenergic receptors: A role in trafficking and mitogen-activated protein kinase activation. Proc Natl Acad Sci U S A 2001;98:343–348. 74. Wilson SJ, Roche AM, Kostetskaia E, Smyth EM. Dimerization of the human receptors for prostacyclin and thromboxane facilitates thromboxane receptor-mediated cAMP generation. J Biol Chem 2004;279:53036–53047. 75. Filipek S, Krzysko KA, Fotiadis D, et al. A concept for G protein activation by G protein-coupled receptor dimers: the transducin/rhodopsin interface. Photochem Photobiol Sci 2004;3:628–638. 76. Lohr HR, Kuntchithapautham K, Sharma AK, Rohrer B. Multiple, parallel cellular suicide mechanisms participate in photoreceptor cell death. Exp Eye Res 2006;83:380–389. 77. Huang P, Gaitan A, Hao Y, Petters R, Wong F. Cellular interactions implicated in the mechanism of photoreceptor degeneration in transgenic mice expressing a mutant rhodopsin gene. Proc Natl Acad Sci U S A 1993;90:8484–8488. 78. Berson EL. Retinitis pigmentosa: unfolding its mystery. Proc Natl Acad Sci U S A 1996;93:4526–4528. 79. Tao YX. Inactivating mutations of G protein-coupled receptors and diseases: structure-function insights and therapeutic implications. Pharmacol Ther 2006;111:949–973.

194

Gleim and Hwa

80. Barak LS, Oakley RH, Laporte SA, Caron MG. Constitutive arrestin-mediated desensitization of a human vasopressin receptor mutant associated with nephrogenic diabetes insipidus. Proc Natl Acad Sci U S A 2001;98:93–98. 81. Min KC, Zvyaga TA, Cypess AM, Sakmar TP. Characterization of mutant rhodopsins responsible for autosomal dominant retinitis pigmentosa. Mutations on the cytoplasmic surface affect transducin activation. J Biol Chem 1993;268:9400–9404. 82. Chuang J-Z, Vega C, Jun W, Sung C-H. Structural and functional impairment of endocytic pathways by retinitis pigmentosa mutant rhodopsin-arrestin complexes. J Clin Invest 2004;114:131–140. 83. Andres A, Kosoy A, Garriga P, Manyosa J. Mutations at position 125 in transmembrane helix III of rhodopsin affect the structure and signalling of the receptor. Eur J Biochem 2001;268:5696–5704. 84. Robinson PR, Cohen GB, Zhukovsky EA, Oprian DD. Constitutively active mutants of rhodopsin. Neuron 1992;9:719–725. 85. Richards JE, Kuo CY, Boehnke M, Sieving PA. Rhodopsin Thr58Arg mutation in a family with autosomal dominant retinitis pigmentosa. Ophthalmology 1991;98:1797–1805. 86. Li T, Franson W, Gordon J, Berson E, Dryja T. Constitutive activation of phototransduction by K296E opsin is not a cause of photoreceptor degeneration. Proc Natl Acad Sci U S A 1995;92:3551–3555. 87. Rao VR, Cohen GB, Oprian DD. Rhodopsin mutation G90D and a molecular mechanism for congenital night blindness. 1994;367:639–642. 88. Najma al-Jandal GJF, Kiang AS, Humphries MM, Bannon N, Findlay JBC, Humphries P, Kenna PF. A novel mutation within the rhodopsin gene (Thr-94-Ile) causing autosomal dominant congenital stationary night blindness. Hum Mutat 1999;13:75–81. 89. Moench SJ, Moreland J, Stewart DH, Dewey TG. Fluorescence studies of the localization and membrane accessibility of the palmitoylation sites of rhodopsin. Biochemistry 1994;33:5791–5796. 90. Moench SJ, Terry CE, Dewey TG. Fluorescence labeling of the palmitoylation sites of rhodopsin. Biochemistry 1994;33:5783–5790. 91. Ernst OP, Meyer CK, Marin EP, et al. Mutation of the fourth cytoplasmic loop of rhodopsin affects binding of transducin and peptides derived from the carboxyl-terminal sequences of transducin alpha and gamma subunits. J Biol Chem 2000;275:1937–1943. 92. Tai AW, Chuang J-Z, Bode C, Wolfrum U, Sung C-H. Rhodopsin’s carboxy-terminal cytoplasmic tail acts as a membrane receptor for cytoplasmic dynein by binding to the dynein light chain Tctex-1. Cell 1999;97:877–887. 93. Garriga P, Liu X, Khorana HG. Structure and function in rhodopsin: Correct folding and misfolding in point mutants at and in proximity to the site of the retinitis pigmentosa mutation Leu-125 right-arrow Arg in the transmembrane helix C. Proc Natl Acad Sci U S A 1996;93:4560–4564. 94. Reeves PJ, Thurmond RL, Khorana HG. Structure and function in rhodopsin: high level expression of a synthetic bovine opsin gene and its mutants in stable mammalian cell lines. Proc Natl Acad Sci U S A 1996;93:11487–11492. 95. Kaushal S, Khorana HG. Structure and function in rhodopsin. 7. Point mutations associated with autosomal dominant retinitis pigmentosa. Biochemistry 1994;33:6121–6128. 96. Doi T, Molday R, Khorana H. Role of the intradiscal domain in rhodopsin assembly and function. Proc Natl Acad Sci U S A 1990;87:4991–4995. 97. Janz JM, Fay JF, Farrens DL. Stability of dark state rhodopsin is mediated by a conserved ion pair in intradiscal loop E-2. J Biol Chem 2003;278:16982–16991.

Rhodopsin and Retinitis Pigmentosa

195

98. Isele J, Sakmar TP, Siebert F. Rhodopsin activation affects the environment of specific neighboring phospholipids: an FTIR spectroscopic study. Biophys J 2000;79:3063–3071. 99. SanGiovanni JP, Chew EY. The role of omega-3 long-chain polyunsaturated fatty acids in health and disease of the retina. Prog Retin Eye Res 2005;24:87–138. 100. Berson EL, Rosner B, Weigel-DiFranco C, Dryja TP, Sandberg MA. Disease progression in patients with dominant retinitis pigmentosa and rhodopsin mutations. Invest Ophthalmol Vis Sci 2002;43:3027–3036. 101. Li T, Sandberg MA, Pawlyk BS, et al. Effect of vitamin A supplementation on rhodopsin mutants threonine-17 right-arrow methionine and proline-347 right-arrow serine in transgenic mice and in cell cultures. Proc Natl Acad Sci U S A 1998;95:11933–11938. 102. Saliba RS, Munro PMG, Luthert PJ, Cheetham ME. The cellular fate of mutant rhodopsin: quality control, degradation and aggresome formation. J Cell Sci 2002;115:2907–2918. 103. Noorwez SM, Kuksa V, Imanishi Y, et al. Pharmacological chaperone-mediated in vivo folding and stabilization of the P23H-opsin mutant associated with autosomal dominant retinitis pigmentosa. J Biol Chem 2003;278:14442–14450. 104. Noorwez SM, Malhotra R, McDowell JH, Smith KA, Krebs MP, Kaushal S. Retinoids assist the cellular folding of the autosomal dominant retinitis pigmentosa opsin mutant P23H. J Biol Chem 2004;279:16278–16284. 105. Kuksa V, Bartl F, Maeda T, et al. Biochemical and physiological properties of rhodopsin regenerated with 11-cis-6-ring- and 7-ring-retinals. J Biol Chem 2002;277:42315–42324. 106. Vogel R, Siebert F, Ludeke S, Hirshfeld A, Sheves M. Agonists and partial agonists of rhodopsin: retinals with ring modifications. Biochemistry 2005;44:11684–11699. 107. Morrison SA, Russell RM, Carney EA, Oaks EV. Zinc deficiency: a cause of abnormal dark adaptation in cirrhotics. Am J Clin Nutr 1978;31:276–281. 108. Dorea JG, Olson JA. The rate of rhodopsin regeneration in the bleached eyes of zinc-deficient rats in the dark. J Nutr 1986;116:121–127. 109. Leure-duPree AE, McClain CJ. The effect of severe zinc deficiency on the morphology of the rat retinal pigment epithelium. Invest Ophthalmol Vis Sci 1982;23:425–434. 110. Kim HJ, Kim YY, Kim SY. Effect of zinc on the visual sensitivity of the bullfrog’s eye. Korean J Ophthalmol 2000;14:53–59. 111. Tam SW, Wilber KE, Wagner FW. Light sensitive zinc content of protein fractions from bovine rod outer segments. Biochem Biophys Res Commun 1976;72:302–309. 112. Shuster TA, Nagy AK, Conly DC, Farber DB. Direct zinc binding to purified rhodopsin and disc membranes. Biochem J 1992;282(Pt 1):123–128. 113. Stojanovic A, Stitham J, Hwa J. Critical role of transmembrane segment zinc binding in the structure and function of rhodopsin. J Biol Chem 2004;279:35932–35941. 114. del Valle LJ, Ramon E, Canavate X, Dias P, Garriga P. Zinc-induced decrease of the thermal stability and regeneration of rhodopsin. J Biol Chem 2003;278:4719–4724. 115. Patel AB, Crocker E, Reeves PJ, et al. Changes in interhelical hydrogen bonding upon rhodopsin activation. J Mol Biol 2005;347:803–812. 116. Swaminath G, T.W. Lee, and B. Kobilka. Identification of an allosteric binding site for Zn2+ on the beta2 adrenergic receptor. J Biol Chem 2003;278:352–356. 117. Swaminath G, et al. Allosteric modulation of beta2-adrenergic receptor by Zn(2+). Mol Pharmacol 2002;61:65–72. 118. Schetz JA, Sibley DR. Zinc allosterically modulates antagonist binding to cloned D1 and D2 dopamine receptors. J Neurochem 1997;68:1990–1997. 119. Holst B, Schwartz TW. Molecular mechanism of agonism and inverse agonism in the melanocortin receptors: Zn(2+) as a structural and functional probe. Ann N Y Acad Sci 2003;994:1–11.

196

Gleim and Hwa

120. Holst B, Elling CE, Schwartz TW. Metal ion-mediated agonism and agonist enhancement in melanocortin MC1 and MC4 receptors. J Biol Chem 2002;277:47662–47670. 121. Wang J, Luthey-Schulten ZA, Suslick KS. Is the olfactory receptor a metalloprotein? Proc Natl Acad Sci U S A 2003;100:3035–3039. 122. Bastek J, Bogden J, Cinotti A, et al. Trace metals in a family with sex-linked retinitis pigmentosa. Adv Exp Med Biol 1977;77:43–50. 123. Mao W, Zeng L, Ma Q, et al. [Primary study of iontophoresis of zinc ion in treatment of retinitis pigmentosa]. Yan Ke Xue Bao 1990;6:88–90. 124. Karcioglu ZA, Stout R, Hahn HJ. Serum zinc levels in retinitis pigmentosa. Curr Eye Res 1984;3:1043–1048.

9 Multiple Signaling Pathways Govern Calcium Homeostasis in Photoreceptor Inner Segments Tamas Szikra and David Krizaj CONTENTS Introduction Overview of Ca2+ Regulation in the Inner Segment Voltage-Operated Calcium Channels Play a Central Role in Inner Segment Calcium Regulation Neurotransmission from Rods and Cones to Second-Order Retinal Neurons Photoreceptor Malfunction and Degeneration Development References

INTRODUCTION The great British physiologist Sydney Ringer was the first to suggest that calcium (Ca2+) plays a central role in coordinating function of excitable cells (Carafoli, 2002). We now know that Ca2+ promotes, modulates, and integrates intracellular signals in all eukaryotic cells by taking advantage of an approximately 10,000-fold driving force for Ca2+ entry into the cytosol. In primary sensory neurons such as photoreceptors, Ca2+ regulates both input (sensory transduction) and output (synaptic transmission), participating as well in additional processes crucial to survival, signaling function, and cell death. This includes regulation of cell growth and development, gene expression, synthesis and release of neurotransmitters, cytoskeletal dynamics, and energy metabolism. This review discusses the cellular mechanisms in vertebrate rods and cones by which Ca2+ manages to carry out this monumental task. The anatomy of vertebrate photoreceptors is similar to that of other primary sensory neurons in that the cells are constructed of two separate anatomical compartments that process signal inputs and outputs, respectively (Fig. 1). An outer segment (OS) is exclusively

From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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Fig. 1. A Generic rod photoreceptor. The outer segment (OS) hosts the phototransduction apparatus. The inner segment (IS) downstream from the OS is formed by three anatomically distinct domains: (1) ellipsoid, which contains most of cell’s mitochondria; (2) the cell body, which contains the cell nucleus, nuclear envelope formed by the endoplasmic reticulum (ER) cisternae; and (3) the synaptic terminal, packed with synaptic vesicles and cisternae of smooth ER. B Dissociated salamander rod and C salamander cone photoreceptor. Ca2+ sequestration and release from the mitochondria occur via Ca2+ uniporter channels and Na+/Ca2+ transporters, respectively. PMCA plasma membrane Ca adenosine triphosphatase (ATPase), VGCC voltage-gated channel. Scale bars 5 µm.

dedicated to transducing photon energy into graded changes in the photoreceptor membrane potential. In contrast, photoreceptor regions downstream from the OS are responsible for life support and survival and for synaptic transmission of light-evoked changes in membrane potential to the rest of the visual system. The inner segment (IS) itself is comprised of a number of anatomically discrete domains (the ellipsoid, the myoid, the cell body, and the synaptic terminal), each of which contains distinct sets of intracellular organelles and ion transporters (Fig. 1). The IS and the OS are connected by incessant large-scale movement of proteins and lipids guided by specialized motors and Ca2+ buffers through a thin nonmotile cilium (Besharse et al., 2003; Giessl et al., 2006) as well as by cyclical light-dependent translocation of at least three phototransduction proteins between the OS and the IS (McGinnis et al., 2002; Sokolov et al., 2002; Strissel et al., 2005). While photoreceptors do not possess a bona fide action potential-conducting axon (their individual neurite may better be thought of as an enlargement of the initial segment of the axon of a conventional neuron), each neurite ends in a specialized synaptic terminal (pedicle in cones, spherule

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Fig. 2. A Simultaneous [Ca2+]i measurement from the cell body and outer segment of a salamander rod. The ryanodine receptor agonist caffeine transiently elevated [Ca2+]i in the inner, but not the outer, segment. B Rod light responses to 200-ms, 567-nm flash (upper panel −3.5 log; bottom panel −1.0 log quanta) before and during caffeine exposure. No significant effect of caffeine was observed on either the transient hyperpolarization of the rod or the rod “tail” for brighter flashes; slightly slower rise times were observed for caffeine exposures during dim flashes. IS inner segment, OS outer segment.

in rods; Lasansky, 1973; Haverkamp et al., 2000). Terminals in mammalian species such as mouse and rat possess a single large mitochondrion, whereas those of amphibian and reptilian photoreceptors appear to lack mitochondria (Lasansky, 1973; Choi et al., 2005). The main point of our chapter is to review how the highly compartmentalized anatomy of rod and cone ISs is associated with region-specific regulation of intracellular calcium concentration [Ca2+] and to relate those differences to homeostatic Ca2+ mechanisms particular to each photoreceptor region. In that regard, the OS and IS express different sets of plasma membrane, intracellular store transporters, and ion channels, which in turn impart differential voltage sensitivity, Ca2+ affinities, and transport and modulation properties to each segment (Krizaj and Copenhagen, 2002). An example is illustrated in Fig. 2, which shows that stimulation of ryanodine receptors evokes large-scale Ca2+ release from internal stores in the rod IS but has no effect on [Ca2+]i homeostasis in the OS or the light response of rod photoreceptors. A great deal has been learned from molecular, physiological, and genetic studies of Ca2+ regulation in the OS (Lamb and Pugh, 2006). These studies established that the OS possesses a single Ca2+ entry pathway (the cyclic guanosine monophosphate [cGMP]dependent [cyclic nucleotide-gated, CNG] channel) and one Ca2+ clearance pathway, the Na,K+/Ca2+ exchanger (NKCX) (Korenbrot and Rebrik, 2002; Palczewski et al., 2004; Paillart et al., 2006). While in some retinas OSs are also labeled by antibodies raised against ryanodine, inositol triphosphate (IP3), and sarcoplasmic-endoplasmic reticulum Ca2+ ATPase (SERCA) channels and transporters (Wang et al., 1999; Krizaj, 2005b; Shoshan-Barmatz et al., 2005), a functional role for specific Ca2+ store transporters in OS Ca2+ regulation still needs to be defined. In contrast to the OS, the IS is characterized by a multitude of Ca2+ influx, clearance, and storage mechanisms, including powerful intracellular organelles such as the endoplasmic reticulum (ER) and mitochondria.

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OVERVIEW OF CA2+ REGULATION IN THE INNER SEGMENT In sensory neurons, Ca2+ acts globally (across tens of micrometers) as well as locally (within “microdomains”) to coordinate, integrate, and tune sensory transduction, gene expression, and neurotransmission through a wide array of signaling systems with differing Ca2+ affinities (e.g., Roberts, 1994; Neher, 1995; Zufall and Leinders-Zufall, 2000; Nakatani et al., 2002a and 2002b; Berridge et al., 2003). Vertebrate rod and cone photoreceptors exemplify this strategy; most, if not all, key biochemical processes in these cells are modulated by spatial, temporal, and amplitude aspects of changes in [Ca2+]i microdomains (reviewed in Fain et al., 2001; Krizaj and Copenhagen, 2002; Heidelberger et al., 2005). [Ca2+]IS is determined by the interplay between activation of Ca2+ entry and clearance mechanisms in the plasma membrane, ER, and mitochondria that keeps [Ca2+]i in darkness high (at 300–600 nM) and low (30–50 nM) in the light (Krizaj et al., 2003; Szikra and Krizaj, 2006). Figure 3 illustrates the pathways for Ca2+ entry into the IS cytosol: (1) plasma membrane voltage-activated Ca2+ channels (Corey et al., 1984); (2) store-operated transient receptor potential canonical (TRPC) or receptor-operated transient receptor potential vanilloid (TRPV)-like channels (Zimov and Yazulla, 2004; Szikra et al., 2006); (3) CNG channels (Rieke and Schwartz, 1994); (4) ryanodine and IP3 receptor-operated release channels (Peng et al., 1991; Krizaj et al., 2004); and (5) Ca2+ release from the mitochondria (Krizaj et al., 2003; Szikra and Krizaj 2006).

Fig. 3. Schema of Ca regulation in the inner segment (IS). The endoplasmic reticulum (ER) Ca store represents a central hub for intracellular Ca homeostasis, communicating with both plasma membrane and mitochondria. Black arrows calcium fluxes, gray arrows activation pathways. CNGC cyclic guanosine monophosphate (cGMP)-gated channel, GPCR G proteincoupled receptor, IP3R, triphosphate (IP3) receptor, NCX mitochondrial Na/Ca exchanger, PLC phospholipase C, PMCA plasma membrane Ca ATPase, RyR ryanodine receptor, SERCA, sarcoplasmic-endoplasmic reticulum Ca adenosine triphosphatase (ATPase), SOC store-operated Ca channel, VOCC L-type voltage-operated Ca channel, CICR, Calcium-Induced Calcium release; DAG, diacyl glycerol; Gprot, G protein.

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Fig. 4. Spatiotemporal [Ca2+]i differences in inner segment (IS) subregions in response to depolarization. A Cone photoreceptor stimulated with 64-ms puffs of KCl. Fast kinetics, high amplitude, and spatially localized depolarization-evoked [Ca2+]i are first observed in the synaptic terminal, followed by the rest of the IS. Note the large spontaneous [Ca2+]i increase in the terminal (arrowhead). B Detail of [Ca2+]i elevation in a rod terminal triggered by transient depolarization. High-amplitude hot spot [Ca2+] increases are observed during depolarization (arrowheads).

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The properties of Ca2+ clearance mechanisms in the IS determine the speed with which photoreceptor output responds to light (Duncan et al., 2006). Mechanisms that clear Ca2+ from IS cytosol consist of (1) plasma membrane Ca2+ adenosine triphosphatase (ATPase) transporters (PMCAs) (Krizaj et al., 2002; Duncan et al., 2006); (2) SERCA transporters (Krizaj et al., 2003, 2004; Krizaj, 2005b); and (3) mitochondrial Ca2+ uniporters (Krizaj et al., 2003) (Fig. 3). Because both PMCAs and SERCAs are driven by metabolic energy in the form of adenosine triphosphate (ATP), Ca2+ homeostasis in photoreceptor ISs is closely linked to energy metabolism derived from glycolytic and oxidative phosphorylation pathways. Steady-state [Ca2+]i amplitude and spatiotemporal characteristics of Ca2+ signals maintained by primary Ca2+ influx and clearance mechanisms are additionally shaped by (1) Ca2+-mediated Ca2+ channel inactivation (Corey et al., 1984; Rabl and Thoreson, 2002); (2) powerful Ca2+-activated chloride and potassium conductances (Barnes and Hille, 1989; Cia et al., 2005; Xu and Slaughter, 2005); (3) glutamate transporter-mediated chloride conductances (Rabl et al., 2003); (4) Ca2+ buffering proteins such as calcium-binding protein (CaBP), parvalbumin, recoverin, and calbindin (Sokal et al., 2000; Haeseleer et al., 2004); (5) retrograde feedback from horizontal cells (Verweij et al., 1996, 2003; Hirasawa and Kaneko, 2003; Vessey et al., 2005); (6) protons released from synaptic vesicles or via metabolism (Kleinschmidt, 1991); (7) Ca2+/H+ exchange (Krizaj and Copenhagen, 1998); (8) chloride modulation (Thoreson et al., 2003); or (9) surface charge (Piccolino et al., 1999; Cadetti et al., 2004). Ca2+-mediated signaling cascades are either confined to specific domains of the IS or are expressed at different densities across several IS regions. As a result, the amplitude and frequency modulation of light-evoked [Ca2+]i levels is highly region specific, allowing for tuning a wide array of Ca2+ signaling systems that use sensors with differing affinities located in different IS domains. For example, voltage-gated Ca2+ channels (VGCCs) and PMCAs are strongly expressed in the terminal and weakly in the ellipsoid region (Rieke and Schwartz, 1996; Nachman-Clewner et al., 1999; Morgans et al., 1998; Krizaj et al., 2002), resulting in large-amplitude, fast [Ca2+] onset and offset kinetics in the terminal (Fig. 4). Ellipsoid [Ca2+] levels are typically at least threefold lower than in the synaptic terminal due to low density of L-type VGCCs and mitochondrial Ca2+ uptake (Szikra and Krizaj, 2006). Ca2+ imaging studies suggested that there may be significant diffusion between the terminal and the cell body as well as between the cell body and the ellipsoid regions (Krizaj and Copenhagen, 1998; Steele et al., 2005; Szikra and Krizaj, 2006). VOLTAGE-OPERATED CALCIUM CHANNELS PLAY A CENTRAL ROLE IN INNER SEGMENT CALCIUM REGULATION The VGCCs represent the central element of Ca2+ regulation in the IS because they possess biophysical properties conferring susceptibility to voltage, heteromeric G proteins, Ca2+ released from stores, and Ca2+ buffers such as calmodulin. The VGCC is a multisubunit protein complex consisting of the principal α1 subunit (which forms the Ca2+ channel pore and senses the membrane potential) and auxiliary β− and α2δ-subunits. In mammalian rods, the α1-subunit belongs to the Cav1.4 (Cav1.4a1q; α1F) subtype transcribed from the L-type family CACNA1F gene (Bech-Hansen et al., 1998). Currents

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through Cav1.4 channels are detected only when α1F subunits are coexpressed with α2δ1− and β2/β3-subunits, indicating that subunit interactions are critical for normal Ca2+ entry into the rod IS (Koschak et al., 2003; Baumann et al., 2004). Deletion of the β2-subunit (but not other β-subunits) alters the IS expression of α1F, resulting in loss of neurotransmission (Ball and Gregg, 2002; but see McRory et al., 2004). Visual deficits in patients with X-linked congenital stationary night blindness (CSNB2; Strom et al., 1998; BechHansen et al., 1998) are caused by mutations in the CACNA1F gene. The mutations underlying the CSNB2 phenotype encompass at least 75 variations. Of these mutations, 23 result in truncated α1-subunits and loss of function; in another 25 cases, various missense mutations were reported in which there still may be some residual function (Hoda et al., 2006). In some mutations, no change in gating was observed; rather, the expression of Cav1.4 was affected (McRory et al., 2004; Hoda et al., 2006), suggesting deficits in protein misfolding or trafficking. Cav1.4 knockout mice are characterized by decreased depolarization-evoked Ca2+ entry into photoreceptor terminals and failure to establish proper targeting/anchoring of presynaptic ribbon complexes in rods (Mansergh et al., 2005). Surprisingly, however, loss of the α1F-subunit resulted in total suppression of both rod and cone neurotransmission (Mansergh et al., 2005), challenging the belief that Cav1.4 are selectively localized to rods. In a recently described gain-of-function phenotype similar to CSNB2, there may be an increase in Ca2+ entry into rod ISs with similarly deleterious effects on the phenotype (Hemara-Wahanui et al., 2005). A number of functional questions need to be addressed with respect to molecular interactions between IS VGCCs and cytosolic signals, extracellular modulators and the membrane potential. To maintain normal function in tonically depolarized cells such as photoreceptors, it is imperative to tame the positive feedback inherent in VGCC activation. A regenerative cycle between Ca2+ influx and depolarization would impede the light response by breaking the relationship between light stimuli and [Ca2+]i; it also would trigger oscillations of about 3–4 Hz in the membrane potential and potentially overload the IS with Ca2+ (Akopian et al., 1997). Photoreceptor ISs employ several strategies to achieve this aim. First, the physiologically relevant range of the I–V relationship is shifted toward negative potentials, minimizing the contribution of Ca2+ current to the steady-state “dark” membrane potential. Hence, in darkness, Ca2+ influx through VGCCs is small (Heidelberger et al., 2005) and amply compensated by counterion fluxes. The α1F-subunit-containing channels in vivo activate at substantially more negative potentials than the large majority of other “high-voltage-activated” Ca2+ channels (Cox and Dunlap, 1992; but see Awatramani et al., 2005). Mammalian photoreceptor VGCC currents activate at −60 mV (primate: Yagi and MacLeish, 1994; tree shrew: Morgans et al., 1998; pig: Cia et al., 2005). Amphibian VGCC currents are first measurable at about −45 mV (Corey et al., 1984; Barnes and Hille, 1989); however, it remains to be determined whether small depolarizations to −60 mV increase [Ca2+]IS in amphibian photoreceptors without evoking detectable Ca2+ currents (Awatramani et al., 2005; Akopian et al., 1997; Bader et al., 1982). Effective voltages measured in vivo are 10–15 mV more negative than α1F-currents observed in heterologously expressing HEK cells (Baumann et al., 2004; McRory et al., 2004), suggesting that the voltage sensitivity and other gating characteristics of indigenous Ca2+ channels are modulated by cytosolic modulators. One such modulator is CaBP4, which shifts the activation curve of the

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Cav1.4 channel to hyperpolarized potentials (Haeseleer et al., 2004). The significance of this modulation is indicated by the finding that loss of CaBP4 results in a 100-fold reduction of sensitivity of mouse rod and cone neurotransmission without affecting OS Ca2+ dynamics (Haeseleer et al., 2004; Maeda et al., 2005). The I–V relationship of IS VGCCs is also modulated by calmodulin, protons, negative feedback from postsynaptic cells, and metabotropic glutamate modulation (Barnes et al., 1993; Verweij et al., 1996; Hosoi et al., 2005; Higgs and Lukasiewicz, 2002). Second, the membrane potential is stabilized at the threshold of VGCC activation through shunts provided by concomitant activation of powerful Ca2+-activated potassium and chloride conductances (Corey et al., 1984; Maricq and Korenbrot, 1988; Barnes and Hille, 1989; Xu and Slaughter, 2005). The delayed rectifier K+ current (reversal potential ~ −80 mV; Beech and Barnes, 1989) and Cl− (reversal potential near −20 mV; Thoreson et al., 2002) counterion fluxes are instrumental for shunting the membrane potential to prevent Ca2+ spiking (Akopian et al., 1997) and Na+ spiking (Kawai et al., 2005; Ohkuma et al., 2006) in rods and cones. Activation of chloride fluxes may also provide negative feedback from postsynaptic horizontal cells (e.g., Verweij et al., 1996, 2003) and directly modulate VGCCs (Thoreson et al., 2003). Third, given that most L-type channels typically experience strong Ca2+-mediated inactivation, how is it possible for IS VGCCs to reliably transduce voltage changes into [Ca2+]i during prolonged depolarizations? All VGCC α1-subunits possess a conserved IQ motif that binds calmodulin, resulting in Ca2+-dependent inactivation due to Ca2+ entering the cytoplasm via channels themselves (Imredy and Yue, 1992), Ca2+ release from the ER (Adachi-Akahane et al., 1996), or release of Ca2+ from mitochondria (Hernandez-Guijo et al., 2001). Recent molecular and physiological studies found that α1F-subunits are much less susceptible to inactivation than α1-subunits in other types of L channels (Baumann et al., 2004; McRory et al., 2004), possibly because of a gating modulator within the C-terminal tail downstream of the IQ domain that regulates calmodulin binding (A. Singh et al., 2006). This inactivation mechanism is important for normal vision because men carrying the truncated IQ domain develop CSNB2 (A. Singh et al., 2006). Whereas little Ca2+-mediated inactivation is seen in heterologously expressed Cav1.4 (Koschak et al., 2003; Baumann et al., 2004; A. Singh et al., 2006), Ca2+-dependent inactivation is more prominent, if still modest, in intact cells (Corey et al., 1984; Rabl et al., 2003). This suggests that, in vivo, α1F inactivation is modulated by cytosolic components within the IS. Voltage-operated Ca2+ entry into ISs is additionally regulated by kinases and phosphatases (Stella et al., 2001, 2002; Akopian et al., 2000; Zhang et al., 2005), protons (Barnes et al., 1993; Hirasawa and Kaneko, 2003), nitric oxide (NO; Kourennyi et al., 2004) and neuromodulators such as dopamine (D2 and D4 receptors), somatostatin (SSA2 receptors), ATP (P2X7 receptors), adenosine (A2 receptors), GABA (γ-aminobutyric acid), insulin, glutamate transporters, and glutamate itself (mGluR group III receptors) that act through heteromeric Gi, Go, and possibly Gq proteins (Stella et al., 2001, 2002; Thoreson et al., 2002; Akopian et al., 2000; Krizaj and Witkovsky, 1993, Higgs and Lukasiewicz, 2002; Koulen et al., 2005; Hosoi et al., 2005; Tatsukawa et al., 2005; Wersinger et al., 2006). Other potential candidates for VGCC modulation are presynaptic proteins such as bassoon (Khimich et al., 2005; tom Dieck et al., 2005), neurexins that link Ca2+ channels to presynaptic PDZ proteins (Missler et al., 2003) and neuroligins

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and syndecans (Song et al., 1999). Finally, the high density of actin in all domains of the IS (Cristofanilli and Akopian, 2006) together with the susceptibility of retinal actin networks to Ca2+ modulation (Akopian et al., 2006) strongly suggest that Ca2+ signaling in the IS is an integral element of developing, remodeling, and regenerating ISs (e.g., Nachman-Clewner et al., 1999). The fascinating panoply of factors that have an impact on or are impacted by [Ca2+]i regulation is a direct indicator of the central role for this second messenger in IS function. VGCCs that drive Ca2+ entry into the IS are selectively targeted to the synaptic terminal, with lesser expression in the cell body and the ellipsoid, as seen in mammalian (Raviola and Gilula, 1975; Morgans et al., 1998; Cia et al., 2005), reptilian (Schaeffer and Raviola, 1976; Lasater and Witkovsky, 1991; Choi et al., 2005), avian (Firth et al., 2001), and amphibian (Nachman-Clewner et al., 1999; Steele et al., 2005; Szikra and Krizaj, 2006) photoreceptors. The targeting mechanisms are unknown. However, VGCCs themselves are known to have a key role in the formation and function of the synapse. Loss of Cav 1.4 α1- or β2-subunits disrupted targeting of the synaptic ribbon to the active zone, suppressed exocytosis, and caused an absence of dendritic extensions of postsynaptic horizontal and bipolar cells into invaginations within the terminal (Ball and Gregg, 2002; Mansergh et al., 2005). This indicates that Ca2+ channels themselves regulate the assembly and trafficking of the synaptic ribbon to the photoreceptor active zone. Finally, the dendrites of postsynaptic horizontal cells mediate retrograde synaptic interactions that control presynaptic [Ca2+]i (Verweij et al., 1996, 2003). Comparison of Boltzman functions fitted to Ca channel activation curves suggested that protonation of VGCCs exerts control over the gain of synaptic transfer by shifting the Ca current–voltage relationship (Barnes et al., 1993; Barnes, 1994; Hirasawa and Kaneko, 2003). The exquisite multiplicity of Ca2+ regulatory machinery organized around VGCCs shows that signal transmission at rod and cone synapses is highly modulatable. Voltage-operated Ca2+ channels in amphibian and mammalian photoreceptors as well as heterologously expressed Cav1.4 channels are about sixfold potentiated by BayK 8644 and inhibited by dihydropyridine blockers such as nifedipine, isradipine, and nimodipine (Wilkinson and Barnes, 1996; Baumann et al., 2004; Szikra and Krizaj, 2006) and by the phenylkylamine verapamil (Cia et al., 2005). These characteristics confirm that CaV1.4 channels are members of the L-type VGCC family. However, sensitivity for dihydropyridine inhibition is uncharacteristically low (Hart et al., 2003; Cia et al., 2005), and some IS Ca2+ channels are reportedly susceptible to N-type Ca2+ channel antagonists such as ω-conotoxin (Wilkinson and Barnes, 1996; Kourennyi and Barnes, 2000). Unlike most L-type channels, heterologously expressed mammalian Cav1.4 subunits (Baumann et al., 2004) and endogenous mammalian VGCCs (Cia et al., 2005) are blocked by both L- and D-cis diltiazem enantiomers. This observation has potent therapeutic implications as diltiazems protect photoreceptors against Ca2+ overload and degeneration (e.g., Frasson et al., 1999; Takano et al., 2004; Vallazza-Deschamps et al., 2005). Ca2+ Channels in Rods and Cones There is a lot of evidence that rods and cones possess different mechanisms of Ca2+ influx and clearance, differential expression of modulatory receptors. as well as different intracellular cascades involving Ca2+ and other second messengers (Krizaj and Witkovsky,

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Fig. 5. Store-operated Ca2+ channels in photoreceptors. Depletion of Ca2+ stores caused a large decrease in baseline [Ca2+]i, followed by a rebound overshoot, a diagnostic marker for storeoperated Ca2+ channels. The overshoot was blocked by the store-operated channel antagonist SKF 96365.

1993; Stella and Thoreson, 2000; Krizaj, 2000; Kourennyi et al., 2004; Zhang et al., 2005; Hosoi et al., 2005; Duncan et al., 2006). These differences are discussed at greater length in Krizaj and Copenhagen (2002). Three examples of compounds with opposite effects on rod and cone VGCCs involve dopamine (Krizaj and Witkovsky, 1993), adenosine (Stella et al., 2002), NO (Kourennyi et al., 2004), and glutamate (Hosoi et al., 2005). Depolarization-induced increases of calcium concentration in rods and cones are enhanced and inhibited, respectively, by the NO donor S-nitrosocysteine (Kourennyi et al., 2004). In contrast, glutamate acting through mGluR receptors suppresses VGCCs in amphibian cones but not rods (Hosoi et al., 2005). Finally, Ca2+ regulation in rods, but not cones, is characterized by large-scale Ca2+ release from ryanodine-based ER stores (Krizaj et al., 2003; Cadetti et al., 2006) (Fig. 5). Hence, rod and cone signals not only possess different timing and spectral sensitivity of the light response in the OS (that arise from expression of rod-specific and cone-specific phototransduction cascade elements), but also have a series of Ca2+ channels, transporters, and buffering systems in the IS that modulate, modify, and shape the final photoreceptor output, reflected in the rate of exocytosis in the synaptic terminal. NEUROTRANSMISSION FROM RODS AND CONES TO SECOND-ORDER RETINAL NEURONS The IS represents a signaling bottleneck in which Ca2+ channels and transporters compress visual stimuli from the 4–5 log response range generated by the phototransduction cascade in the OS to about 0.5 log dynamic range of presynaptic [Ca2+]IS (Szikra and Krizaj, 2006) and transmitter release (Choi et al., 2005). The compression reflects a cumulative superposition of numerous positive- and negative-feedback loops inherent in biophysical and neuromodulatory properties of channels and transporters. Thus, transmission of graded light-evoked signals across the photoreceptor synapse is controlled and modulated by voltage-activated and Ca2+-activated mechanisms within the IS. The exquisite accuracy of neurotransmission at photoreceptor synapses is preserved by the sensitivity of graded transmitter release to changes in presynaptic [Ca2+]i.

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This release appears to be linearly proportional to [Ca2+]IS over the full range of presynaptic membrane potentials (Rieke and Schwartz, 1996; Witkovsky et al., 1997; Thoreson et al., 2004). The linearity of the relationship between Ca2+ influx and transmitter release in photoreceptors is difficult to understand because Ca2+ action in presynaptic terminals tends to be highly cooperative (Dodge and Rahamimoff, 1967; Goda and Stevens, 1994; Bollmann et al., 2000). It is possible that linearity is caused by linear summation of independent Ca2+ entry sites (e.g., Heidelberger et al., 2005). Alternatively, the linearity may result from linear activation of Ca2+ extrusion mechanisms during the light response because the PMCA pumps that regulate the light response are activated proportionally to [Ca2+]i (Zador and Koch, 1994; Duncan et al., 2006). Finally, the linear operating range of presynaptic [Ca2+]i might be sculpted by release of Ca2+ from intracellular stores (Cadetti et al., 2006) or Ca2+ entry through store-operated Ca2+permeable channels (Szikra et al., 2006). A series of recent findings showed that Ca2+ release from intracellular stores plays a crucial role in rod neurotransmission (Krizaj et al., 1999; Cadetti et al., 2006; Suryanarayanan and Slaughter, 2006). The predominant role of Calcium-Induced Calcium Release (CICR) at rod synapses is unparalleled in the central nervous system. CICR represents an amplification mechanism that boosts exocytosis in the darkness, when photoreceptors are depolarized. Consistent with this, CICR inhibitors most affect glutamate release during stimulation with dim flashes (Suryanarayanan and Slaughter, 2006), and synaptic gain is steepest around the dark membrane potential (see Witkovsky et al., 1997; Belgum and Copenhagen, 1988). The magnitude of Ca2+ release from stores is set by the “basal” cytosolic [Ca2+]i (Krizaj et al., 1999) as Ca2+; entering the IS through VGCCs triggers further release of Ca2+ from the ER by activating ryanodine receptor channels within the ER (Krizaj et al., 2003). Ryanodine receptors belong to the cardiac RyR isoform 2 family (Krizaj et al., 2004). CICR contributes most to synaptic [Ca2+]i during long depolarizations with Ca2+ fluxes that cross the threshold for ryanodine receptor activation. At these potentials, global elevation of [Ca2+]i measured in synaptic terminals (400–2000 nM; Szikra and Krizaj, 2006) triggers exocytosis at rates ranging from about 250 vesicles/s in lizard cones to about 400 vesicles/s in amphibian rods (Rieke and Schwartz, 1996; Schmitz and Witkovsky, 1997; Choi et al., 2005). The capacitance method combined with Ca2+ imaging was used to measure exocytosis directly in isolated photoreceptors. The threshold for exocytosis when [Ca2+]i was measured with high-affinity indicators fura-2, fura-FF, and fluo-3 was under 1 µM (Rieke and Schwartz, 1996; Thoreson et al., 2004). However, fura-2 cannot be used to estimate the actual Ca2+ concentration required for transmitter release as [Ca2+]i close to sites of Ca2+ entry can approach tens to hundreds of micromoles, well above the about 2 µM saturation for fura-2 (Neher, 1995; Sinha et al., 1997; Llinas et al., 1992a,b). Studies using the low-affinity Ca2+ indicator furaptra at ribbon synapses suggested that exocytosis at ribbon synapses has a threshold of about 10 µM (photoreceptors; Kreft et al., 2003) or 50 µM (teleost bipolar cells; Heidelberger et al., 1994), consistent with the idea that vesicle release is determined by local [Ca2+]i microdomains. Similar low- and high-affinity mechanisms were reported in bipolar cells following use of low- and highaffinity indicators, respectively (Heidelberger et al., 1994; von Gersdorff and Matthews, 1996; Zhou et al., 2006).

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Alternatively, exocytosis in photoreceptors may be regulated by high- and low-affinity Ca2+ sensors. Photoreceptor terminals express synaptotagmin I, a relatively low-affinity (Kd 3–30 µM free Ca2+) presynaptic “release sensor” through which primed vesicles within the terminal detect high peaks of localized Ca2+ (Geppert and Sudhof, 1998) (cow: von Kriegstein et al., 1999; salamander: Kreft et al., 2003; mouse: Lazzell et al., 2004). The higher affinity isoform, synaptotagmin III, was recently identified in rat, mouse, and goldfish photoreceptors (Berntson and Morgans, 2003). It is possible that synaptotagmin I drives the fast synchronous component, whereas synaptotagmin III drives the slower, asynchronous component, possibly in cooperation with synaptotagmin XII, a Ca2+independent regulator of release (e.g., Maximov and Sudhof, 2005; Maximov et al., 2007). Finally, a fast exocytotic component could be driven by Ca2+ influx through VGCCs, whereas asynchronous release might be more susceptible to CICR (Suryanarayanan and Slaughter, 2006). Consistent with these ideas, kinetics of exocytosis in rod photoreceptors exhibits fast and slow components, with vesicle fusing to the plasma membrane with rate constants of about 2 and about 127 ms, respectively (Kreft et al., 2003). Blocking CICR blocked the slow, but not fast, component of postsynaptic excitatory postsynaptic current (EPSC) in bipolar cells (Suryanarayanan and Slaughter, 2006). Neuromodulators strongly affect photoreceptor neurotransmission. As noted, presynaptic [Ca2+] signals (and neurotransmitter glutamate release) are modulated by many agents, including dopamine, somatostatin, ATP, adenosine, insulin, NO, and glutamate itself. For example, group II mGluR agonists inhibit glutamate release from salamander rods and cones (Higgs and Lukasiewicz, 2002), whereas group III agonists were effective in mouse rods (Koulen et al., 2005) and newt cones (Hosoi et al., 2005). PHOTORECEPTOR MALFUNCTION AND DEGENERATION By using calcium ions as an intracellular messenger, cells walk a tightrope between life and death. Too much or too little Ca2+ at the wrong time and place might lead to rapid cell death by necrosis or to the induction of the cell death program of apoptosis (Paschen, 2003). Typically, main features of the apoptotic process include activation of the caspase protease superfamily, release of proapoptic factors such as cytochrome c from the ER and mitochondria, and a pathological increase in cytosolic [Ca2+] (Ca2+ overload) that causes membrane blebbing, fragmentation of nucleosomal DNA, mitochondrial malfunction, and cell death (Hajnoczky et al., 2003; Mattson and Chan, 2003). Pathological increases in [Ca2+]i have been implicated in a number of degenerative and agent-evoked photoreceptor diseases, including retinitis pigmentosa (Chang et al., 1993), lead-induced toxicity models (He et al., 2000; Fox et al., 2003), hypoxia (Toescu, 2004), retinal detachment (Khodair et al., 2003; Zhang et al., 2005), and cancer-associated retinopathy (Subramanian and Polans, 2004; Adamus et al., 2006). Ca2+ overload is an early and rapid event in photoreceptor degeneration induced by prolonged exposure to light (Donovan et al., 2001, Fox et al., 2003; Doonan et al., 2005; Wenzel et al., 2005). Increased levels of photoreceptor [Ca2+]i have been measured immediately after toxic light exposure, with the number of rods with elevated [Ca2+]i continuing to increase up to 3 h (Donovan and Cotter, 2002; Donovan et al., 2001). In degenerative diseases, Ca2+ overloads increase gradually as cells enter the apoptotic process during the development

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(Doonan et al., 2005). Impaired calcium regulation in the IS has also been implicated in diseases such as Duchenne muscular dystrophy (Fitzgerald et al., 1994) and CSNB2 (Bech-Hansen et al., 1998; Hemara-Wahanui et al., 2005). Correspondingly, when Ca2+ overload is prevented by a genetic deletion of VGCC subunits (Read et al., 2002) or by using pharmacological antagonists of Ca2+ channels (Edward et al., 1991; Takano et al., 2004), photoreceptor cell death is reduced. Ca2+ overload is pathological for cells because it is typically followed by secondary insults that involve activation of multidestructive pathways such as calcineurin phosphatases, cathepsin, and calpain proteases and inflammation, together with activation of “conventional” apoptotic signals (consisting of Bax/Bid signaling, the caspase system, increased production of reactive oxygen species, and excessive Ca2+ loading of mitochondria; Hajnoczky et al., 2003, 2006; Mattson and Chan, 2003). While the apoptotic process in photoreceptors sometimes appears to be independent of mitochondrial release of cytochrome c or caspase activation observed in inner retinal cells (Cusato et al., 2003; Donovan and Cotter, 2002), this is not always the case. A major cause of blindness in humans is mutations in genes that encode proteins of the phototransduction cascade. Mutations in phosphodiesterase β subunit (PDEβ6), guanylate cyclase (GC), and guanylate cyclase-activating protein (GCAP) (Sokal et al., 2000; Palczewski et al., 2004; L. Jiang et al., 2005) lead to elevated [cGMP]OS and Ca2+ influx through CNG channels (Chang et al., 1993; Farber 1995), which ultimately results in activation of the apoptotic program. Increases in [cGMP] were directly shown to cause Ca2+ overload in the rd1 mouse (a model for autosomal recessive human retinitis pigmentosa) and in degenerating amphibian photoreceptors (Lolley et al., 1977; Lolley, 1994; Vallazza-Deschamps et al., 2005). In mouse photoreceptors, intracellular [Ca2+] levels increased just before cells entered apoptosis at P10 (Doonan et al., 2005). This cytosolic increase was associated with mitochondrial depolarization, suggesting that degenerating photoreceptors experience a concomitant mitochondrial Ca2+ overload (e.g., Fox et al., 2003). Interestingly, some forms of photoreceptor degeneration may be caused by a decrease in [Ca2+]OS (Fain, 2006). Consistent with this, suppression of CNG channels with antisense RNA was reported to cause photoreceptor degeneration (Leconte and Barnstable, 2000). How Ca2+ overload generated by excessive activation of CNG channels in the OS is communicated to apoptotic pathways in the IS is not known. Intracellular stores are the essential loci for induction of apoptosis in eukaryotic cells. Overloading and redistribution of Ca2+ within the ER and mitochondria amplify apoptotic signals, and Ca2+ release from these organelles directly activates transcriptional cascades regulating the apoptotic pathway (Breckenridge et al., 2003; Scorrano et al., 2003). A number of recent studies suggested that intracellular organelles contribute to apoptotic signals in rods and cones. Mitochondria, damaged by heavy metals such as lead, were suggested to kill rods through Ca2+ overload mediated by activation of the permeability transition pore in the inner membrane (He et al., 2003; Fox et al., 2003). Suppression of Ca2+ sequestration into ER kills differentiating photoreceptors (Linden, 1999; Chiarini et al., 2003), presumably due to global suppression of mRNA translation that occurs in cells with depleted Ca2+ stores (Brostrom and Brostrom, 2003). Consistent with this idea, intra-ER Ca2+ buffers such as calnexin enhance cell survival by preventing

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store depletion and cytosolic Ca2+ overload (Rosenbaum et al., 2006). Rod photoreceptor cell death induced by store depletion is suppressed by inhibitors of protein synthesis (Chiarini et al., 2003), further underscoring the importance of ER Ca2+ regulation for photoreceptor survival. Finally, direct links between ER Ca content and the caspase cascade were uncovered by the finding that the proapoptotic effect of store depletion is blocked by the pan-caspase inhibitor z-VAD-fmk (Chiarini et al., 2003). Proapoptotic Ca2+ overload can also occur as a result of Ca2+ entry via store-operated or transient receptor potential (TRP) channels (Toescu, 2004). In invertebrate photoreceptors, pathological Ca2+ entry through TRP channels overloads cells’ mitochondria and triggers cell degeneration and death (Yoon et al., 2000; Minke and Agam, 2003). TRP proteins are prominently expressed in rod and cone photoreceptors (Zimov and Yazulla, 2004; Rohrer and Krizaj, in preparation), suggesting that store-operated Ca2+ entry occurs in these cells (e.g., Fig. 5) (Szikra and Krizaj, 2006; Szikra et al., 2006). There is increasing evidence that Ca2+ overload triggers activation of calpains, a family of ER cysteine proteases with substrates that include cytoskeletal proteins such as fodrin as well as proapoptotic proteins Bax, p35, p53, procaspase 9, procaspase 3, and poly ADP ribose polymerase (PARP) (Nicotera and Orrenius, 1998; Rohrer et al., 2004, Paquet-Durand, Azadi et al., 2006). Calpains appear to be positioned at the epicenter of photoreceptor degeneration. Increases in calpain-mediated proteolysis in rods exposed to light-evoked degeneration protocols were observed within 30 min of light induction and continued to increase up to 24 h after the onset of light (Donovan and Cotter, 2002). Calpain activation was also reported in light-independent rd1 degeneration models (Sharma and Rohrer, 2004), in rd1 retinas (Doonan et al., 2005), and in retinas from RCS (Royal College of Surgeons) rats (another model for inherited retinal degeneration; Azarian and Williams, 1995). Activation of calpain-mediated proteolysis consists of two steps that result in translocation of activated calpain from the ER into cytosol. First, the endogenous inhibitor calpastatin must be separated from the calpain heterodimer, causing its translocation from ER lumen to the cytosolic side of the ER. Second, subsequent Ca2+ binding leads to a conformational change and dissociation of the 28-kDa regulatory subunit from the 80-kDa catalytic subunit (Hood et al., 2004). A publication suggested that increased calpain activation in rd1 retinas (Doonan et al., 2005) is not caused by increased calpain transcription or expression per se but rather by its increased dissociation from calpastatin within ER stores (Pacquet-Durand et al., 2006). It remains to be seen whether calpain inhibitors ameliorate photoreceptor degeneration, as seen in other tissues (Ray et al., 1999; DeBiasi et al., 2001). Perhaps to compensate, long- term increases in expression of genes coding for calcium-binding proteins such as calbindin has been observed in rd1 retinas (Hackam et al., 2004; Rohrer et al., 2004). Pathological increases in [Ca2+]i could trigger apoptosis through activation of conventional apoptotic signals involving caspase cascades. The exact involvement of caspases in photoreceptor degeneration seems to depend on the species and the type of injury/disease. The cytosolic pathway involving caspases 9 and 3 is upregulated in the rd model (Sharma and Rohrer, 2004), lead toxicity (He et al., 2000), tubby mice (Bode and Wolfrum, 2003), and cultured cells expressing properties of cone photoreceptors that were stressed by serum withholding (Doonan et al., 2005) but not in

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light-induced photoreceptor degeneration (Donovan et al., 2001; Donovan and Cotter, 2002). Caspase 1, but not caspase 3, may be activated during ischemia (Katai and Yoshimura, 1999) and light-induced degeneration (Samardzija et al., 2006); loss of caspase 1 correlates with the photoreceptor’s ability to withstand cell death induced by excessive light exposure or ischemic injury (Arai et al., 2006) but does not prevent its rapid death in the rd1 degeneration model (Samardzija et al., 2006). Apoptosis in some photoreceptor death models is blocked by the pan-caspase inhibitor z-VAD-fmk (Chiarini et al., 2003; Doonan et al., 2005; Duenker et al., 2005), whereas in other models caspase inhibitors failed to show significant neuroprotection (Donovan and Cotter, 2002; Yoshizawa et al., 2002; Zeiss et al., 2004; Gomez-Vicente et al., 2005; see also Lohr et al., 2006). Caspases and calpains share many substrates, resulting in cross-activation of their respective signals (Berliocchi et al., 2005). For example, caspases facilitate calpain activity through cleavage of calpastatin (Pörn-Ares et al., 1998). Conversely, calpains directly cleave caspases, activating them as well as proapoptotic Bcl-2 family members that reside in intracellular stores. Such parallel activation of calpains and caspases was detected in cultured cone photoreceptors (Doonan et al., 2005). Finally, apoptotic and nonapoptotic death signals can be intertwined. In addition to activation by Ca2+, caspases can also cause pathological Ca2+ overload by inactivating PMCAs (Leist et al., 1998). Relatively little photoreceptor death is observed in most types of CSNB2, which also is associated with scant vascular and RPE changes (usually seen in degenerative eye diseases) (Bech-Hansen et al., 1998). As opposed to loss of photoreceptor nuclei in rods lacking the β2-subunit (Ball et al., 2002) or in retinitis pigmentosa, there is no retinal degeneration in CSNB2 animal models or human patients. The phenotype in this disease, caused by pathological expression of voltage-activated Ca2+ channels in rod ISs, is thus functional rather than anatomical. While light responses of CSNB2 rods are normal, the significant decreases in the amplitude of b-wave electroretinograms in human CSNB2 patients reveal a defect in neurotransmission between rod photoreceptors and bipolar cells (Miyake et al., 1986; Tremblay et al., 1995; Mansergh et al., 2005). Now, we know this is due to loss or mutation in the CACNA1F gene and its Cav1.4 Ca channel product (Bech-Hansen et al., 1998; see above). Therapeutic Strategies The pharmacological strategies for treating retinal degenerations have mainly been based on attempts to suppress apoptosis mediated by Ca overload or caspases. A number of studies reported successful outcomes. For example, calpain inhibition can attenuate ischemic retinal damage (Sakamoto et al., 2000; Gilgun-Sherki et al., 2002; Osborne et al., 2002). Moreover, light-induced degeneration could be blocked with D-cis diltiazem (Doonan et al., 2003; Frasson et al., 1999; Yamazaki et al., 2002; Takano et al., 2004), whereas D-cis diltiazem enhanced photoreceptor survival in the rd mouse and RCS rat (Frasson et al., 1999; Vallazza-Deschamps et al., 2005; but see Bush et al., 2000; Pawlyk et al., 2002; Pearce-Kelling et al., 2001). Consistent with original results by Frasson et al. (1999), deletion of L-type VGCCs from rd1 photoreceptors slowed the progression of degeneration (Read et al., 2002). In the RCS model, the preservation mediated by the L-type channel antagonist nilvadipine was associated with an upregulation

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of neurotrophic factors such as fibroblast growth factor 2 (FGF-2; Sato et al., 2003) and downregulation of caspases 1 and 2 (Yamazaki et al., 2002). L-cis Diltiazem, but not L-type channel blockers verapamil and D-cis diltiazem, was effective in ameliorating heavy-metal toxic effects in rods (Fox et al., 2003). DEVELOPMENT Spontaneous calcium transients are a central feature of the developing retina, and they have well-defined roles in the development of the visual system (Syed et al., 2004; reviewed in Firth et al., 2005; Metea and Newman, 2006). These include gap-junctional Ca2+ wave communication between retinal progenitors (Pearson et al., 2004) involving L-type VGCCs, Ca2+ stores, and store-operated Ca2+-permeable channels (Firth et al., 2005; Sugioka et al., 1999). Ca2+ transients are directly correlated with interkinetic movement of neuroblast nuclei along the radial glia, a movement that is reduced by 81% by BAPTA-AM and by blockers of gap junctions (Pearson et al., 2005). Photoreceptor development is associated with specific Ca2+ signaling pathways (such as PMCAs and SERCAs; Rentería et al., 2005; Krizaj, 2005b) and with specific activation patterns of heterotrimeric G proteins and gap junctions (muscarinic receptors and purinergic P2Y receptors; Pearson et al., 2002). Ca2+ stores play an essential role in development, possibly regulating DNA synthesis transitions between S-, G1, and G2 phases (Sugioka and Yamashita, 2003). It will be interesting to see to what extent Ca2+-mediated photoreceptor degeneration pathways delineated here recapitulate activation of dormant early development pathways. Large-scale apoptosis occurring during normal photoreceptor development (particularly during the onset of cytogenesis and synaptogenesis) and in degenerating rods and cones is associated with large Ca2+ fluxes (Mervin and Stone, 2002). It is possible that developing rods and cones require greater Ca2+ fluxes and higher [Ca2+]i than their mature counterparts to facilitate survival, synapse formation, and other cellular functions (e.g., Rentería et al., 2005; Ghosh and Greenberg, 1995). In conclusion, the diverse and complex repertoire of Ca2+ signals in the IS serves to integrate sensory transduction with essential biological systems needed for cell survival, metabolism, gene expression, translation, and control of apoptosis. This review highlights some of the Ca2+-dependent mechanisms and pathways that are required to sustain photoreceptor and retinal function. ACKNOWLEDGMENTS Tamas Szikra is presently at the Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland. We thank Prof. Paul Witkovsky (New York School of Medicine) for comments on the manuscript. The work was supported by a grant from the Hungarian Eötvös Fellowship and the Knights Templar Foundation (T.S.), by the National Institutes of Health (EY13870), and That Man May See Foundation to D.K. and an unrestricted grant from Research to Prevent Blindness to the University of Utah’s Department of Ophthalmology. Also, Dr. Krizaj is a recipient of a Research to Prevent Blindness James S. Adams Award.

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REFERENCES Adachi-Akahane S, Cleemann L, Morad M. Cross-signaling between L-type Ca2+ channels and ryanodine receptors in rat ventricular myocytes. J Gen Physiol. 1996;108:435–454. Adamus G, Webb S, Shiraga S and Duvoisin SM. Anti-recoverin antibodies induce an increase in intracellular calcium, leading to apoptosis in retinal cells. J Autoimmun. 2006;26:146–153. Akopian A, Johnson J, Gabriel R, Brecha N, Witkovsky P. Somatostatin modulates voltage-gated K(+) and Ca(2+) currents in rod and cone photoreceptors of the salamander retina. J Neurosci. 2000;20:929–936. Akopian A, Krizaj D, Witkovsky P. Both high- and low voltage-activated calcium currents contribute to the light-evoked responses of luminosity horizontal cells in the Xenopus retina. Brain Res. 1997;762:121–130. Akopian A, Szikra T, Cristofanilli M, Krizaj D. Glutamate-induced Ca2+ influx in third-order neurons of salamander retina is regulated by the actin cytoskeleton. Neuroscience. 2006;138:17–24. Arai J, Katai N, Kuida K, Kikuchi T, Yoshimura N. Decreased retinal neuronal cell death in caspase-1 knockout mice. Jpn J Ophthalmol. 2006;50:417–425. Awatramani GB, Price GD, Trussell LO. Modulation of transmitter release by presynaptic resting potential and background calcium levels. Neuron. 2005;48:109–121. Azarian SM, Williams DS. Calpain activity in the retinas of normal and RCS rats. Curr Eye Res. 1995;14:731–735. Bader CR, Bertrand D, Schwartz EA. Voltage-activated and calcium-activated currents studied in solitary rod inner segments from the salamander retina. J Physiol. 1982;331:2532–2584. Baldridge WH, Kurennyi DE, Barnes S. Calcium-sensitive calcium influx in photoreceptor inner segments. J Neurophysiol. 1998;79:3012–3018. Ball SL, Gregg RG. Using mutant mice to study the role of voltage-gated calcium channels in the retina. Adv Exp Med Biol. 2002;514:439–450. Barnes S. After transduction: response shaping and control of transmission by ion channels of the photoreceptor inner segments. Neuroscience. 1994;58:447–459. Barnes S, B. Hille: Ionic channels of the inner segment of tiger salamander cone photoreceptors. J Gen Physiol. 1989;94:719–743. Barnes S, Merchant V, Mahmud FM. Modulation of transmission gain by protons at the photoreceptor output synapse. Proc Natl Acad Sci U S A. 1993;90:10081–10085. Baumann L, Gerstner A, Zong X, Biel M, Wahl-Schott C. Functional characterization of the L-type Ca2+ channel Cav1.4alpha1 from mouse retina. Invest Ophthalmol Vis Sci. 2004;45:708–713. Bech-Hansen NT, Naylor MJ, Maybaum TA, Pearce WG, Koop B, Fishman GA, Mets M, Musarella MA, Boycott KM. Loss-of-function mutations in a calcium-channel alpha1subunit gene in Xp11.23 cause incomplete X-linked congenital stationary night blindness. Nat Genet. 1998;19:264–267. Beech DJ, Barnes S. Characterization of a voltage-gated K+ channel that accelerates the rod response to dim light. Neuron. 1989;3:573–581. Belgum JH, Copenhagen DR. Synaptic transfer of rod signals to horizontal and bipolar cells in the retina of the toad (Bufo marinus). J Physiol. 1988;396:225–245. Berliocchi L, Fava E, Leist M, Horvat V, Dinsdale D, Read D, Nicotera P. Botulinum neurotoxin C initiates two different programs for neurite degeneration and neuronal apoptosis. J Cell Biol. 2005;168:607–618. Berntson AK, Morgans CW. Distribution of the presynaptic calcium sensors, synaptotagmin I/II and synaptotagmin III, in the goldfish and rodent retinas. J Vis. 2003;3:274–280. Berridge MJ, Bootman MD, Roderick HL. Calcium signalling: dynamics, homeostasis and remodelling. Nat Rev Mol Cell Biol. 2003;4:517–529.

214

Szikra and Krizaj

Besharse JC, Baker SA, Luby-Phelps K, Pazour GJ. Photoreceptor intersegmental transport and retinal degeneration: a conserved pathway common to motile and sensory cilia. Adv Exp Med Biol. 2003;533:157–164. Bode C, Wolfrum U. Caspase-3 inhibitor reduces apototic photoreceptor cell death during inherited retinal degeneration in tubby mice. Mol Vis. 2003;9:144–150. Bollmann JH, Sakmann B, Borst JG. Calcium sensitivity of glutamate release in a calyx-type terminal. Science. 2000;289:953–957. Breckenridge DG, Germain M, Mathai JP, Nguyen M, Shore GC. Regulation of apoptosis by endoplasmic reticulum pathways. Oncogene. 2003;22:8608–8618. Brostrom MA, Brostrom CO. Calcium dynamics and endoplasmic reticular function in the regulation of protein synthesis: implications for cell growth and adaptability. Cell Calcium. 2003;34:345–363. Bush RA, Kononen L, Machida S, Sieving PA. The effect of calcium channel blocker diltiazem on photoreceptor degeneration in the rhodopsin Pro213His rat. Invest Ophthalmol Vis Sci. 2000;41:2697–2701. Cadetti L, Bryson EJ, Ciccone CA, Rabl K, Thoreson WB. Calcium-induced calcium release in rod photoreceptor terminals boosts synaptic transmission during maintained depolarization. Eur J Neurosci. 2006;23:2983–2990. Cadetti L, Thoreson WB, Piccolino M. Pre- and post-synaptic effects of manipulating surface charge with divalent cations at the photoreceptor synapse. Neuroscience. 2004;129: 791–801. Carafoli E. Calcium signaling: a tale for all seasons. Proc Natl Acad Sci U S A. 2002;99:1115–1122. Chang GQ, Hao Y, Wong F. Apoptosis: final common pathway of photoreceptor death in rd, rds, and rhodopsin mutant mice. Neuron. 1993;11:595–605. Chiarini LB, Leal-Ferreira ML, de Freitas FG, Linden R. Changing sensitivity to cell death during development of retinal photoreceptors. J Neurosci Res 2003;74:875–883. Choi SY, Borghuis BG, Rea R, Levitan ES, Sterling P, Kramer RH. Encoding light intensity by the cone photoreceptor synapse. Neuron. 2005;48:555–562. Cia D, Bordais A, Varela C, Forster V, Sahel JA, Rendon A, Picaud S. Voltage-gated channels and calcium homeostasis in mammalian rod photoreceptors. J Neurophysiol. 2005;93:1468–1475. Corey DP, Dubinsky JM, Schwartz EA. The calcium current in inner segments of rods from the salamander (Ambystoma tigrinum) retina. J Physiol 1984;354:557–575. Cox DH, Dunlap K. Inactivation of N-type calcium current in chick sensory neurons: calcium and voltage dependence. J Gen Physiol. 1994;104:311–336. Cristofanilli M, Akopian A. Calcium channel and glutamate receptor activities regulate actin organization in salamander retinal neurons. J Physiol. 2006;575:543–554. Cusato K, Bosco A, Rozental R, Guimaraes CA, Reese BE, Linden R, Spray DC. Gap junctions mediate bystander cell death in developing retina. J Neurosci. 2003;23:6413–6422. DeBiasi RL, Edelstein CL, Sherry B, Tyler KL. Calpain inhibition protects against virus-induced apoptotic myocardial injury. J Virol. 2001;75:351–361. Dodge FA Jr, Rahamimoff R. Co-operative action a calcium ions in transmitter release at the neuromuscular junction. J Physiol. 1967;193:419–432. Donovan M, Carmody RJ, Cotter TG. Light-induced photoreceptor apoptosis in vivo requires neuronal nitric-oxide synthase and guanylate cyclase activity and is caspase-3-independent. J Biol Chem. 2001;276:23000–23008. Donovan M, Cotter TG. Caspase-independent photoreceptor apoptosis in vivo and differential expression of apoptotic protease activating factor-1 and caspase-3 during retinal development. Cell Death Differ. 2002;9:1220–1231.

Ca2+ Regulation in the Photoreceptor Inner Segment

215

Doonan F, Donovan M, Cotter TG. Caspase-independent photoreceptor apoptosis in mouse models of retinal degeneration. J Neurosci. 2003;23:5723–5731. Doonan F, Donovan M, Cotter TG. Activation of multiple pathways during photoreceptor apoptosis in the rd mouse. Invest Ophthalmol Vis Sci. 2005;46:3530–3538. Duenker N, Valenciano AI, Franke A, Hernandez-Sanchez C, Dressel R, Behrendt M, De Pablo F, Krieglstein K, de la Rosa EJ. Balance of pro-apoptotic transforming growth factor-beta and anti-apoptotic insulin effects in the control of cell death in the postnatal mouse retina. Eur J Neurosci. 2005;22:28–38. Duncan J, Yang H, Doan T, Silverstein B, Murphy G., Nune G, Liu X, Copenhagen DR, Rieke F, Tempel BL, Krizaj D. Scotopic visual signaling in the mouse retina is modulated by high affinity plasma membrane calcium extrusion J Neurosci. 2006;26:7201–7211. Edward DP, Lam TT, Shahinfar S, Li J, Tso MO. Amelioration of light-induced retinal degeneration by a calcium overload blocker. Flunarizine. Arch Ophthalmol. 1991;109:554–562. Fain GL, Matthews HR, Cornwall MC, Koutalos Y. Adaptation in vertebrate photoreceptors. Physiol Rev. 2001;81:117–151. Fain GL. Why photoreceptors die (and why they don’t). Bioessays. 2006;28:344–354. Farber DB. From mice to men: the cyclic GMP phosphodiesterase gene in vision and disease. Invest Ophthalmol Vis Sci. 1995;36:263–275. Firth SI, Morgan IG, Boeken MK, Morgans C. Localization of voltage-sensitive L-type calcium channels in the chicken retina. Clin Experiment Ophthalmol. 2001;29:183–187. Firth SI, Wang CT, Feller MB. Retinal waves: mechanisms and function in visual system development. Cell Calcium. 2005;37:425–432. Fitzgerald KM, Cibis GW, Giambrone SA, Harris DJ. Retinal signal transmission in Duchenne muscular dystrophy: evidence for dysfunction in photoreceptor/depolarizing bipolar cell pathway. J Clin Invest. 1994;93:2425–2430. Fox DA, Poblenz AT, He L, Harris JB, Medrano CJ. Pharmacological strategies to block rod photoreceptor apoptosis caused by calcium overload: a mechanistic target-site approach to neuroprotection. Eur J Ophthalmol. 2003;3:S44–S56. Frasson M, Sahel JA, Fabre M, Simonutti M, Dreyfus H, Picaud S. Retinitis pigmentosa: rod photoreceptor rescue by a calcium-channel blocker in the rd mouse. Nat Med. 1999;5:1183–1187. Geppert M, Sudhof TC. RAB3 and synaptotagmin: the yin and yang of synaptic membrane fusion. Annu Rev Neurosci. 1998;21:75–95. Ghosh A, Greenberg ME. Calcium signaling in neurons: molecular mechanisms and cellular consequences. Science. 1995;268:239–247. Review. Giessl A, Trojan P, Rausch S, Pulvermuller A, Wolfrum U. Centrins, gatekeepers for the lightdependent translocation of transducin through the photoreceptor cell connecting cilium. Vision Res. 2006;46:4502–4509. Gilgun-Sherki Y, Rosenbaum Z, Melamed E, Offen D. Antioxidant therapy in acute central nervous system injury: current state. Pharmacol Rev. 2002;54:271–384. Goda Y, Stevens CF. Two components of transmitter release at a central synapse. Proc Natl Acad Sci U S A. 1994;91:12942–12946. Gomez-Vicente V, Donovan M, Cotter TG. Multiple death pathways in retina-derived 661 W cells following growth factor deprivation: crosstalk between caspases and calpains. Cell Death Differ. 2005;12:796–804. Hackam AS, Strom R, Liu D, Qian J, Wang C, Otteson D, Gunatilaka T, Farkas RH, Chowers I, Kageyama M, Leveillard T, Sahel JA, Campochiaro PA, Parmigiani G, Zack DJ. Identification of gene expression changes associated with the progression of retinal degeneration in the rd1 mouse. Invest Ophthalmol Vis Sci. 2004;45:2929–2942.

216

Szikra and Krizaj

Haeseleer F, Imanishi Y, Maeda T, Possin DE, Maeda A, Lee A, Rieke F, Palczewski K. Essential role of Ca2+-binding protein 4, a Cav1.4 channel regulator, in photoreceptor synaptic function. Nat Neurosci. 2004;7:1079–1087. Hajnoczky G, Csordas G, Das S, Garcia-Perez C, Saotome M, Sinha Roy S, Yi M. Mitochondrial calcium signalling and cell death: approaches for assessing the role of mitochondrial Ca2+ uptake in apoptosis. Cell Calcium. 2006;40:553–560. Review. Hajnoczky G, Davies E, Madesh M. Calcium signaling and apoptosis. Biochem Biophys Res Commun. 2003;304:445–454. Hart J, Wilkinson MF, Kelly ME, Barnes S. Inhibitory action of diltiazem on voltage-gated calcium channels in cone photoreceptors. Exp Eye Res. 2003;76:597–604. Haverkamp S, Grunert U, Wassle H. The cone pedicle, a complex synapse in the retina. Neuron. 2000;27:85–95. He L, Perkins GA, Poblenz AT, Harris JB, Hung M, Ellisman MH, Fox DA. Bcl-xL overexpression blocks bax-mediated mitochondrial contact site formation and apoptosis in rod photoreceptors of lead-exposed mice. Proc Natl Acad Sci U S A. 2003;100:1022–1027. He L, Poblenz AT, Medrano CJ, Fox DA. Lead and calcium produce rod photoreceptor cell apoptosis by opening the mitochondrial permeability transition pore. J Biol Chem. 2000;275:12175–12184. Heidelberger R, Heinemann C, Neher E, Matthews G. Calcium dependence of the rate of exocytosis in a synaptic terminal. Nature. 1994;371:513–515. Heidelberger R, Thoreson WB, Witkovsky P. Synaptic transmission at retinal ribbon synapses. Prog Retin Eye Res. 2005;24:682–720. Hemara-Wahanui A, Berjukow S, Hope CI, Dearden PK, Wu SB, Wilson-Wheeler J, Sharp DM, Lundon-Treweek P, Clover GM, Hoda JC, Striessnig J, Marksteiner R, Hering S, Maw MA. A CACNA1F mutation identified in an X-linked retinal disorder shifts the voltage dependence of Cav1.4 channel activation. Proc Natl Acad Sci U S A. 2005;102:7553–7558. Hernandez-Guijo JM, Maneu-Flores VE, Ruiz-Nuno A, Villarroya M, Garcia AG, Gandia L. Calcium-dependent inhibition of L, N, and P/Q Ca2+ channels in chromaffin cells: role of mitochondria. J Neurosci. 2001;21:2553–2560. Higgs MH, Lukasiewicz PD. Activation of group II metabotropic glutamate receptors inhibits glutamate release from salamander retinal photoreceptors. Vis Neurosci. 2002;19:275–281. Hirasawa H, Kaneko A. pH changes in the invaginating synaptic cleft mediate feedback from horizontal cells to cone photoreceptors by modulating Ca2+ channels. J Gen Physiol. 2003;122:657–671. Hoda JC, Zaghetto F, Singh A, Koschak A, Striessnig J. Effects of congenital stationary night blindness type 2 mutations R508Q and L1364H on Cav1.4 L-type Ca2+ channel function and expression. J Neurochem. 2006;96:1648–1658. Hood JL, Brooks WH, Roszman TL. Differential compartmentalization of the calpain/calpastatin network with the endoplasmic reticulum and Golgi apparatus. J Biol Chem. 2004;279: 43126–43135. Hosoi N, Arai I, Tachibana M. Group III metabotropic glutamate receptors and exocytosed protons inhibit L-type calcium currents in cones but not in rods. J Neurosci. 2005;25:4062–4072. Imredy JP, Yue DT. Submicroscopic Ca2+ diffusion mediates inhibitory coupling between individual Ca2+ channels. Neuron. 1992;9:197–207. Jiang L, Katz BJ, Yang Z, Zhao Y, Faulkner N, Hu J, Baird J, Baehr W, Creel DJ, Zhang K. Autosomal dominant cone dystrophy caused by a novel mutation in the GCAP1 gene (GUCA1A) Mol Vis. 2005;11:143–151. Katai N, Yoshimura N. Apoptotic retinal neuronal death by ischemia-reperfusion is executed by two distinct caspase family proteases. Invest Ophthalmol Vis Sci. 1999;40:2697–2705.

Ca2+ Regulation in the Photoreceptor Inner Segment

217

Kawai F, Horiguchi M, Ichinose H, Ohkuma M, Isobe R, Miyachi E. Suppression by an h current of spontaneous Na+ action potentials in human cone and rod photoreceptors. Invest Ophthalmol Vis Sci. 2005;46:390–397. Khimich D, Nouvian R, Pujol R, Tom Dieck S, Egner A, Gundelfinger ED, Moser T. Hair cell synaptic ribbons are essential for synchronous auditory signalling. Nature. 2005;434:889–894. Khodair MA, Zarbin MA, Townes-Anderson E. Synaptic plasticity in mammalian photoreceptors prepared as sheets for retinal transplantation. Invest Ophthalmol Vis Sci. 2003;44:4976–4988. Kleinschmidt J. Signal transmission at the photoreceptor synapse. Role of calcium ions and protons. Ann N Y Acad Sci. 1991;635:468–470. Korenbrot JI, Rebrik TI. Tuning outer segment Ca2+ homeostasis to phototransduction in rods and cones. Adv Exp Med Biol. 2002;514:179–203. Koschak A, Reimer D, Walter D, Hoda JC, Heinzle T, Grabner M, Striessnig J. Cav1.4alpha1 subunits can form slowly inactivating dihydropyridine-sensitive L-type Ca2+ channels lacking Ca2+-dependent inactivation. J Neurosci. 2003;23:6041–6049. Koulen P, Liu J, Nixon E, Madry C. Interaction between mGluR8 and calcium channels in photoreceptors is sensitive to pertussis toxin and occurs via G protein betagamma subunit signaling Invest Ophthalmol Vis Sci. 2005;46:287–291. Kourennyi DE, Barnes S. Depolarization-induced calcium channel facilitation in rod photoreceptors is independent of G proteins and phosphorylation. J Neurophysiol. 2000;84:133–138. Kourennyi DE, Liu XD, Hart J, Mahmud F, Baldridge WH, Barnes S. Reciprocal modulation of calcium dynamics at rod and cone photoreceptor synapses by nitric oxide. J Neurophysiol. 2004;92:477–483. Kreft M, Krizaj D, Grilc S, Zorec R. Properties of exocytotic response in vertebrate photoreceptors. J Neurophysiol. 2003;90:218–225. Krizaj D. Mesopic state: cellular mechanisms involved in pre- and post-synaptic mixing of rod and cone signals. Microsc Res Tech. 2000;50:347–359. Review. Krizaj D. Compartmentalization of calcium entry pathways in mouse rods. Eur J Neurosci. 2005a;22:3292–3296. Krizaj D. Serca isoform expression in the mammalian retina. Exp Eye Res. 2005b;81:690–699. Krizaj D, Bao JX, Schmitz Y, Witkovsky P, Copenhagen DR. Caffeine-sensitive calcium stores regulate synaptic transmission from retinal rod photoreceptors. J Neurosci. 1999;19:7249–7261. Krizaj D, Copenhagen DR. Compartmentalization of calcium extrusion mechanisms in the outer and inner segments of photoreceptors. Neuron 1998;21:249–256. Krizaj D, Copenhagen DR. Calcium regulation in photoreceptors. Front Biosci. 2002;7:d2023– d2044. Krizaj D, Demarco SJ, Johnson J, Strehler EE, Copenhagen DR. Cell-specific expression of plasma membrane calcium ATPase isoforms in retinal neurons. J Comp Neurol. 2002;451:1–21. Krizaj D, Lai FA, Copenhagen DR. Ryanodine stores and calcium regulation in the inner segments of salamander rods and cones. J Physiol. 2003;547:761–774. Krizaj D, Liu XL, Copenhagen DR. Expression of calcium transporters in the retina of the tiger salamander (Ambystoma tigrinum). J Comp Neurol. 2004;475:463–480. Krizaj D, Witkovsky P. Effects of submicromolar concentrations of dopamine on photoreceptor to horizontal cell communication. Brain Res. 1993;627:122–128. Lamb TD, Pugh EN Jr. Phototransduction, dark adaptation, and rhodopsin regeneration the proctor lecture. Invest Ophthalmol Vis Sci. 2006;47:5137–5152.

218

Szikra and Krizaj

Lasansky A. Organization of the outer synaptic layer in the retina of the larval tiger salamander. Philos Trans R Soc Lond B Biol Sci. 1973;265:471–489. Lasater EM, Witkovsky P. The calcium current of turtle cone photoreceptor axon terminals. Neurosci Res Suppl. 1991;15:S165–S173. Lazzell DR, Belizaire R, Thakur P, Sherry DM, Janz R. SV2B regulates synaptotagmin 1 by direct interaction. J Biol Chem. 2004;279:52124–52131. Leconte L, Barnstable CJ. Impairment of rod cGMP-gated channel alpha-subunit expression leads to photoreceptor and bipolar cell degeneration. Invest Ophthalmol Vis Sci. 2000;41:917–926. Leist M, Volbracht C, Fava E, Nicotera P. 1-Methyl-4-phenylpyridinium induces autocrine excitotoxicity, protease activation, and neuronal apoptosis. Mol Pharmacol. 1998;54:789–801. Linden R, Rehen SK, Chiarini LB. Apoptosis in developing retinal tissue. Prog Retinal Eye Res. 1999;18:133–165. Llinas R, Sugimori M, Silver RB. Microdomains of high calcium concentration in a presynaptic terminal. Science. 1992a;256:677–679. Llinas R, Sugimori M, Silver RB. Presynaptic calcium concentration microdomains and transmitter release. J Physiol Paris. 1992b;86:135–138. Lohr HR, Kuntchithapautham K, Sharma AK, Rohrer B. Multiple, parallel cellular suicide mechanisms participate in photoreceptor cell death. Exp Eye Res. 2006;83:380–389. Lolley RN. The rd gene defect triggers programmed rod cell death. The Proctor Lecture. Invest Ophthalmol Vis Sci. 1994;35:4182–4191. Lolley RN, Farber DB, Rayborn ME, Hollyfield JG. Cyclic GMP accumulation causes degeneration of photoreceptor cells: simulation of an inherited disease. Science. 1977;196:664–666. Maeda T, Lem J, Palczewski K, Haeseleer F. A critical role of CaBP4 in the cone synapse. Invest Ophthalmol Vis Sci. 2005;46:4320–4327. Mansergh F, Orton NC, Vessey JP, Lalonde MR, Stell WK, Tremblay F, Barnes S, Rancourt DE, Bech-Hansen NT. Mutation of the calcium channel gene Cacna1f disrupts calcium signaling, synaptic transmission and cellular organization in mouse retina. Hum Mol Genet. 2005 Oct 14:3035–3046. Maricq AV, Korenbrot JI. Calcium and calcium-dependent chloride currents generate action potentials in solitary cone photoreceptors. Neuron. 1988;1:503–515. Mattson MP, Chan SL. Calcium orchestrates apoptosis. Nat Cell Biol. 2003;5:1041–1043. Maximov A, Shin OH, Liu X, Sudhof TC. Synaptotagmin-12, a synaptic vesicle phosphoprotein that modulates spontaneous neurotransmitter release. J Cell Biol. 2007;176:113–124. Maximov A, Sudhof TC. Autonomous function of synaptotagmin 1 in triggering synchronous release independent of asynchronous release. Neuron. 2005;48:547–554. McGinnis JF, Matsumoto B, Whelan JP, Cao W. Cytoskeleton participation in subcellular trafficking of signal transduction proteins in rod photoreceptor cells. J Neurosci Res. 2002;67:290–297. McRory JE, Hamid J, Doering CJ, Garcia E, Parker R, Hamming K, Chen L, Hildebrand M, Beedle AM, Feldcamp L, Zamponi GW, Snutch TP. The CACNA1F gene encodes an L-type calcium channel with unique biophysical properties and tissue distribution. J Neurosci. 2004;24:1707–1718. Mervin K, Stone J. Developmental death of photoreceptors in the C57BL/6J mouse: association with retinal function and self-protection. Exp Eye Res. 2002;75:703–713. Metea MR, Newman EA. Calcium signaling in specialized glial cells. Glia. 2006;54:650–655. Minke B, Agam K. TRP gating is linked to the metabolic state and maintenance of the Drosophila photoreceptor cells. Cell Calcium. 2003;33:395–408. Missler M, Zhang W, Rohlmann A, Kattenstroth G, Hammer RE, Gottmann K, Sudhof TC. Alphaneurexins couple Ca2+ channels to synaptic vesicle exocytosis. Nature. 2003;423:939–948.

Ca2+ Regulation in the Photoreceptor Inner Segment

219

Miyake Y, Yagasaki K, Horiguchi M, Kawase Y, Kanda T. Congenital stationary night blindness with negative electroretinogram. A new classification. Arch Ophthalmol. 1986;104:1013–1020. Morgans CW, El Far O, Bernstson A, Wassle H, Taylor WR. Calcium extrusion from mammalian photoreceptor terminals. J Neurosci 1998;18:2467–2474. Nachman-Clewner M, St Jules R, Townes-Anderson E. L-type calcium channels in the photoreceptor ribbon synapse: localization and role in plasticity. J Comp Neurol. 1999;415:1–16. Nakatani K, Chen C, Koutalos Y. Calcium diffusion coefficient in rod photoreceptor outer segments. Biophys J. 2002;82:728–739. Nakatani K, Chen C, Yau KW, Koutalos Y. Calcium and phototransduction. Adv Exp Med Biol. 2002;514:1–20. Neher E. The use of fura-2 for estimating Ca buffers and Ca fluxes. Neuropharmacology. 1995;34:1423–1442. Neher E. Vesicle pools and Ca2+ microdomains: new tools for understanding their roles in neurotransmitter release. Neuron. 1998;20:389–399. Nicotera P, Orrenius S. The role of calcium in apoptosis. Cell Calcium. 1998;23:173–180. Ohkuma M, Kawai F, Horiguchi M, Miyachi EI. Patch-clamp recording of human retinal photoreceptors and bipolar cells. Photochem Photobiol. 2007;83:317–322. Osborne NN, Wood JPM, Cupido A, Melena J, Chidlow G. Topical flunarizine reduces IOP and protects the retina against ischemia-excitotoxicity. Invest Ophthalmol Vis Sci. 2002;43: 1456–1464. Paillart C, Zhang K, Rebrik TI, Baehr W, Korenbrot JI. Cloning and molecular characterization of cGMP-gated ion channels from rod and cone photoreceptors of striped bass (M. saxatilis) retina. Vis Neurosci. 2006;23:99–113. Palczewski K, Sokal I, Baehr W. Guanylate cyclase-activating proteins: structure, function, and diversity. Biochem Biophys Res Commun. 2004;322:1123–1130. Review. Paquet-Durand F, Azadi D, Hauck SM, Uefong M, Veen T, Ekstrom P. Calpain is activated in degenerating photoreceptors in the rd1 mouse. J Neurochem. 2006;96:802–814. Paquet-Durand F, Johnson L, Ekstrom P. Calpain activity in retinal degeneration. J Neurosci Res. 2007;85:693–702. Paschen W. Mechanisms of neuronal cell death: diverse roles of calcium in the various subcellular compartments. Cell Calcium. 2003;34:305–310. Pawlyk BS, Li T, Scimeca MS, Sandberg MA, Berson EL. Absence of photoreceptor rescue with D-cis-diltiazem in the rd mouse. Invest Ophthalmol Vis Sci. 2002;43:1912–1915. Pearce-Kelling SE, Aleman TS, Nickle A, Laties AM, Aguirre GD, Jacobson SG, Acland GM. Calcium channel blocker D-cis-diltiazem does not slow retinal degeneration in the PDE6B mutant rcd1 canine model of retinitis pigmentosa. Mol Vis. 2001;25:42–47. Pearson RA, Catsicas M, Becker DL, Bayley P, Luneborg NL, Mobbs P. Ca(2+) signalling and gap junction coupling within and between pigment epithelium and neural retina in the developing chick. Eur J Neurosci. 2004;19:2435–2445. Pearson R, Catsicas M, Becker D, Mobbs P. Purinergic and muscarinic modulation of the cell cycle and calcium signaling in the chick retinal ventricular zone. J Neurosci. 2002;22:7569–7579. Pearson RA, Luneborg NL, Becker DL, Mobbs P. Gap junctions modulate interkinetic nuclear movement in retinal progenitor cells. J Neurosci. 2005;16:10803–10814. Peng YW, Sharp AH, Snyder SH, Yau KW. Localization of the inositol 1,4,5-trisphosphate receptor in synaptic terminals in the vertebrate retina. Neuron. 1991;6:525–531. Piccolino M, Vellani V, Rakotobe LA, Pignatelli A, Barnes S, McNaughton P. Manipulation of synaptic sign and strength with divalent cations in the vertebrate retina: pushing the limits of tonic, chemical neurotransmission. Eur J Neurosci. 1999;11:4134–4138.

220

Szikra and Krizaj

Pörn-Ares MI, Samali A, Orrenius S. Cleavage of the calpain inhibitor, calpastatin, during apoptosis. Cell Death Differ. 1998;5:1028–1033. Rabl K, Bryson EJ, Thoreson WB. Activation of glutamate transporters in rods inhibits presynaptic calcium currents. Vis Neurosci. 2003;20:557–566. Rabl K, Thoreson WB. Calcium-dependent inactivation and depletion of synaptic cleft calcium ions combine to regulate rod calcium currents under physiological conditions. Eur J Neurosci. 2002;16:2070–2077. Raviola E, Gilula NB. Intramembrane organization of specialized contacts in the outer plexiform layer of the retina. A freeze-fracture study in monkeys and rabbits. J Cell Biol. 1975;65:192–222. Ray SK, Wilford GG, Matzelle DC, Hogan EL, Banik NL. Calpeptin and methylprednisolone inhibit apoptosis in rat spinal cord injury. Ann N Y Acad Sci. 1999;890:261–269. Read DS, McCal MA and Gregg RG. Absence of voltage-dependent calcium channels delays photoreceptor degeneration in rd mice Exp. Eye Res. 2002;75:415–420. Rentería RC, Strehler EE, Copenhagen DR, Krizaj D. Ontogeny of plasma membrane Ca2+ ATPase isoforms in the neural retina of the postnatal rat. Vis Neurosci. 2005;22:263–274. Rieke F, Schwartz EA. A cGMP-gated current can control exocytosis at cone synapses. Neuron. 1994;13:863–873. Rieke F, Schwartz EA. Asynchronous transmitter release: control of exocytosis and endocytosis at the salamander rod synapse. J Physiol. 1996;493:1–8. Roberts WM. Localization of calcium signals by a mobile calcium buffer in frog saccular hair cells. J Neurosci. 1994;14:3246–3262. Rohrer B, Pinto FR, Hulse KE, Lohr HR, Zhang L, Almeida JS. Multidestructive pathways triggered in photoreceptor cell death of the rd mouse as determined through gene expression profiling. J Biol Chem. 2004;279:41903–41910. Rosenbaum EE, Hardie RC, Colley NJ. Calnexin is essential for rhodopsin maturation, Ca2+ regulation, and photoreceptor cell survival. Neuron. 2006;49:229–241. Sakamoto YR, Nakajima TR, Fukiage CR, Sakai OR, Yoshida YR, Azuma MR, Shearer TR. Involvement of calpain isoforms in ischemia-reperfusion injury in rat retina. Curr Eye Res. 2000;21:571–580. Samardzija M, Wenzel A, Thiersch M, Frigg R, Reme C, Grimm C. Caspase-1 ablation protects photoreceptors in a model of autosomal dominant retinitis pigmentosa. Invest Ophthalmol Vis Sci. 2006;47:5181–5190. Sato M, Ohguro H, Ohguro I, Mamiya K, Takano Y, Yamazaki H, Metoki T, Miyagawa Y, Ishikawa F, Nakazawa M. Study of pharmacological effects of nilvadipine on RCS rat retinal degeneration by microarray analysis. Biochem Biophys Res Commun. 2003;306:826–831. Schaeffer SF, Raviola E. Ultrastructural analysis of functional changes in the synaptic endings of turtle cone cells. Cold Spring Harb Symp Quant Biol. 1976;40:521–528. Schmitz Y, Witkovsky P. Dependence of photoreceptor glutamate release on a dihydropyridinesensitive calcium channel. Neuroscience. 1997;78:1209–1216. Scorrano L, Oakes SA, Opferman JT, Cheng EH, Sorcinelli MD, Pozzan T, Korsmeyer SJ. BAX and BAK regulation of endoplasmic reticulum Ca2+: a control point for apoptosis. Science. 2003;300:135–139. Sharma AK, Rohrer B. Calcium-induced calpain mediates apoptosis via caspase-3 in a mouse photoreceptor cell line. J Biol Chem. 2004;279:35564–35572. Shoshan-Barmatz V, Orr I, Martin C, Vardi N. Novel ryanodine-binding properties in mammalian retina. Int J Biochem Cell Biol. 2005;37:1681–1695. Singh A, Hamedinger D, Hoda JC, Gebhart M, Koschak A, Romanin C, Striessnig J. C-terminal modulator controls Ca2+-dependent gating of Ca(v)1.4 L-type Ca2+ channels. Nat Neurosci. 2006;9:1108–1116. Sinha SR, Wu LG, Saggau P. Presynaptic calcium dynamics and transmitter release evoked by single action potentials at mammalian central synapses. Biophys J. 1997;72:637–651.

Ca2+ Regulation in the Photoreceptor Inner Segment

221

Sokal I, Li N, Verlinde CLMJ, Haeseleer F, Baehr W, Palczewski K. Ca-binding proteins in the retina: from discovery to etiology of human disease. Biochim Biophys Acta. 2000;1498: 233–251. Sokolov M, Lyubarsky AL, Strissel KJ, Savchenko AB, Govardovskii VI, Pugh EN Jr, Arshavsky VY. Massive light-driven translocation of transducin between the two major compartments of rod cells: a novel mechanism of light adaptation. Neuron. 2002;34:95–106. Song JY, Ichtchenko K, Sudhof TC, Brose N. Neuroligin 1 is a postsynaptic cell-adhesion molecule of excitatory synapses. Proc Natl Acad Sci U S A. 1999;96:1100–1105. Steele EC, Chen X, Iuvone PM, MacLeish PR. Imaging of Ca2+ dynamics within the presynaptic terminals of salamander rod photoreceptors. J Neurophysiol. 2005;94:4544–4553. Stella SL Jr, Bryson EJ, Thoreson WB. Insulin inhibits voltage-dependent calcium influx into rod photoreceptors. Neuroreport. 2001;12:947–951. Stella SL, Bryson EJ, Thoreson WB. A2 adenosine receptors inhibit calcium influx through L-type calcium channels in rod photoreceptors of the salamander retina. J Neurophysiol. 2002;87:351–360. Stella SL Jr, Thoreson WB. Differential modulation of rod and cone calcium currents in tiger salamander retina by D2 dopamine receptors and cAMP. Eur J Neurosci. 2000;12:3537–3548. Strissel KJ, Lishko PV, Trieu LH, Kennedy MJ, Hurley JB, Arshavsky VY. Recoverin undergoes light-dependent intracellular translocation in rod photoreceptors. J Biol Chem. 2005;280:29250–29255. Strom TM, Nyakatura G, Apfelstedt-Sylla E, Hellebrand H, Lorenz B, Weber BH, Wutz K, Gutwillinger N, Ruther K, Drescher B, Sauer C, Zrenner E, Meitinger T, Rosenthal A, Meindl A. An L-type calcium-channel gene mutated in incomplete X-linked congenital stationary night blindness. Nat Genet. 1998;19:260–263. Subramanian L, Polans AS. Cancer-related diseases of the eye: the role of calcium and calciumbinding proteins. Biochem Biophys Res Commun. 2004;322:1153–1165. Sugioka M, Yamashita M. Calcium signaling to nucleus via store-operated system during cell cycle in retinal neuroepithelium. Neurosci Res. 2003;45:447–458. Sugioka M, Zhou WL, Hofmann HD, Yamashita M. Ca2+ mobilization and capacitative Ca2+ entry regulate DNA synthesis in cultured chick retinal neuroepithelial cells. Int J Dev Neurosci. 1999;17:163–172. Suryanarayanan A, Slaughter MM. Synaptic transmission mediated by internal calcium stores in rod photoreceptors. J Neurosci. 2006;26:1759–1766. Syed MM, Lee S, He S, Zhou ZJ. Spontaneous waves in the ventricular zone of developing mammalian retina. J Neurophysiol. 2004;91:1999–2009. Szikra T, Cusato K, Krizaj D. Store-operated channels regulate baseline Ca in light-adapted photoreceptors. IOVS Abstr 2006, Ft. Lauderdale, FL. Szikra T, Krizaj D. The dynamic range and domain-specific signals of intracellular calcium in photoreceptors. Neuroscience. 2006;141:143–155. Takano Y, Ohguro H, Dezawa M, Ishikawa H, Yamazaki H, Ohguro I, Mamiya K, Metoki T, Ishikawa F, Nakazawa M. Study of drug effects of calcium channel blockers on retinal degeneration of rd mouse. Biochem Biophys Res Commun. 2004;313:1015–1022. Tatsukawa T, Hirasawa H, Kaneko A, Kaneda M. GABA-mediated component in the feedback response of turtle retinal cones. Vis Neurosci. 2005;22:317–324. Thoreson WB, Bryson EJ, Rabl K. Reciprocal interactions between calcium and chloride in rod photoreceptors. J Neurophysiol. 2003;90:1747–1753. Thoreson WB, Rabl K, Townes-Anderson E, Heidelberger R. A highly Ca2+-sensitive pool of vesicles contributes to linearity at the rod photoreceptor ribbon synapse. Neuron. 2004;42:595–605. Thoreson WB, Stella SL Jr, Bryson EI, Clements J, Witkovsky P. D2-like dopamine receptors promote interactions between calcium and chloride channels that diminish rod synaptic transfer in the salamander retina. Vis Neurosci. 2002;19:235–247.

222

Szikra and Krizaj

Toescu EC. Hypoxia sensing and pathways of cytosolic Ca2+ increases. Cell Calcium. 2004;36:187–199. tom Dieck S, Altrock WD, Kessels MM, Qualmann B, Regus H, Brauner D, Fejtová A, Barcko O, Gundelfinger ED, Brandstätter JH. Molecular dissection of the photoreceptor ribbon synapse: physical interaction of Bassoon and RIBEYE is essential for the assembly of the ribbon complex. J Cell Biol. 2005;168:825–836. Tremblay F, Laroche RG, De Becker I. The electroretinographic diagnosis of the incomplete form of congenital stationary night blindness. Vision Res. 1995;35:2383–2393. Vallazza-Deschamps G, Fuchs C, Cia D, Tessier LH, Sahel JA, Dreyfus H, Picaud S. Diltiazeminduced neuroprotection in glutamate excitotoxicity and ischemic insult of retinal neurons. Doc Ophthalmol. 2005;110:25–35. Verweij J, Hornstein EP, Schnapf JL. Surround antagonism in macaque cone photoreceptors. J Neurosci. 2003;23:10249–10257. Verweij J, Kamermans M, Spekreijse H. Horizontal cells feed back to cones by shifting the cone calcium-current activation range. Vision Res. 1996;36:3943–3953. Vessey JP, Stratis AK, Daniels BA, Da Silva N, Jonz MG, Lalonde MR, Baldridge WH, Barnes S. Proton-mediated feedback inhibition of presynaptic calcium channels at the cone photoreceptor synapse. J Neurosci. 2005;25:4108–4117. von Gersdorff H, Matthews G. Calcium-dependent inactivation of calcium current in synaptic terminals of retinal bipolar neurons. J Neurosci. 1996;16:115–122. von Kriegstein K, Schmitz F, Lonk E, Sudhof T. Distribution of synaptic vesicle proteins in the mammalian retina identifies obligatory and facultative components of ribbon synapses. Eur J Neurosci. 1999;11:1335–1348. Wang TL, Sterling P, Vardi N. Localization of type I inositol 1,4,5-triphosphate receptor in the outer segments of mammalian cones. J Neurosci. 1999;19:4221–4228. Wenzel A, Grimm C, Samardzija M, Reme CE. Molecular mechanisms of light-induced photoreceptor apoptosis and neuroprotection for retinal degeneration. Prog Retinal Eye Res. 2005;24:275–306. Wersinger E, Schwab Y, Sahel JA, Rendon A, Pow DV, Picaud S, Roux MJ. The glutamate transporter EAAT5 works as a presynaptic receptor in mouse rod bipolar cells. J Physiol. 2006;577:221–234. Wilkinson MF, Barnes S. The dihydropyridine-sensitive calcium channel subtype in cone photoreceptors. J Gen Physiol. 1996;107:621–630. Witkovsky P, Schmitz Y, Akopian A, Krizaj D, Tranchina D. Gain of rod to horizontal cell synaptic transfer: relation to glutamate release and a dihydropyridine-sensitive calcium current. J Neurosci. 1997;17:7297–7306. Xu JW, Slaughter MM. Large-conductance calcium-activated potassium channels facilitate transmitter release in salamander rod synapse. J Neurosci. 2005;25:7660–7668. Yagi T, MacLeish PR. Ionic conductances of monkey solitary cone inner segments. J Neurophysiol. 1994;71:656–665. Yamazaki H, Ohguro H, Maeda T, Maruyama I, Takano Y, Metoki T, Nakazawa M, Sawada H, Dezawa M. Preservation of retinal morphology and functions in royal college surgeons rat by nilvadipine, a Ca(2+) antagonist. Invest Ophthalmol Vis Sci. 2002;43:919–926. Yoon J, Ben-Ami HC, Hong YS, Park S, Strong LL, Bowman J, Geng C, Baek K, Minke B, Pak WL. Novel mechanism of massive photoreceptor degeneration caused by mutations in the trp gene of Drosophila. J Neurosci. 2000;20:649–659. Yoshizawa K, Kiuchi K, Nambu H, Yang J, Senzaki H, Kiyozuka Y, Shikata N, Tsubura A. Caspase-3 inhibitor transiently delays inherited retinal degeneration in C3H mice carrying the rd gene. Graefes Arch Clin Exp Ophthalmol. 2002;240:214–219.

Ca2+ Regulation in the Photoreceptor Inner Segment

223

Zador A, Koch C. Linearized models of calcium dynamics: formal equivalence to the cable equation. J Neurosci. 1994;14:4705–4715. Zeiss CJ, Neal J, Johnson EA. Caspase-3 in postnatal retinal development and degeneration. Invest Ophthalmol Vis Sci. 2004;45:964–970. Zhang N, Beuve A, Townes-Anderson E. The nitric oxide-cGMP signaling pathway differentially regulates presynaptic structural plasticity in cone and rod cells. J Neurosci. 2005;25:2761–2770. Zhou ZY, Wan QF, Thakur P, Heidelberger R. Capacitance measurements in the mouse rod bipolar cell identify a pool of releasable synaptic vesicles. J Neurophysiol. 2006;96:2539–2548. Zimov S, Yazulla S. Localization of vanilloid receptor 1 (TRPV1/VR1)-like immunoreactivity in goldfish and zebrafish retinas: restriction to photoreceptor synaptic ribbons. J Neurocytol. 2004;33:441–452. Zufall F, Leinders-Zufall T. The cellular and molecular basis of odor adaptation. Chem Senses. 2000;25:473–481.

10 The Transduction Channels of Rod and Cone Photoreceptors Dimitri Tränkner CONTENTS The Transduction Channels of Rod and Cone Photoreceptors The Role of CNG Channels in Photoreceptor Physiology The Molecular Composition of CNG Channels The Basic Activation Properties of CNG Channels Transmembrane Topology and Functional Domains CNG Channels Are Components of Larger Protein Complexes Modulation by Phosphorylation and All-TRANS Retinal Synthesis, Maturation, and Targeting of CNG Channels Visual Dysfunction Caused by Mutant CNG Channel Genes References

THE TRANSDUCTION CHANNELS OF ROD AND CONE PHOTORECEPTORS In contrast to most sensory receptor cells that depolarize in response to a stimulus, vertebrate photoreceptors respond to light with brief hyperpolarization. In darkness, photoreceptors maintain elevated levels of the intracellular messenger guanosine 3′,5′-cyclic monophosphate (cGMP) that directly activate their transduction channels, the so-called cyclic nucleotide-gated (CNG) ion channels. In the dark, active CNG channels conduct a sustained depolarizing inward current carried by sodium (Na+) and calcium (Ca2+) ions. Light triggers a cascade of enzymatic reactions that culminates in the degradation of cGMP, closure of the CNG channels, and photoreceptor hyperpolarization. Besides functioning as cGMP sensors, CNG channels serve a critical role in the light adaptation of photoreceptors. Light adaptation is Ca2+ dependent and therefore responsive to changes in the Ca2+ flux through CNG channels. The pivotal role of CNG channels in photoreceptor physiology is reflected in the fact that the two photoreceptor types, rods and cones, utilize differently tuned CNG channels to meet specific tasks. The following sections provide an overview of the functional aspects and cell biology of CNG channels in photoreceptors in the normal and diseased state. From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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THE ROLE OF CNG CHANNELS IN PHOTORECEPTOR PHYSIOLOGY Visual perception in humans operates over a dynamic range of more than nine orders of magnitude, from the light conditions of an overcast night to bright daylight with glare reflecting from a glacier. This extraordinarily broad range is made possible by two different types of photoreceptors. Rods, with an exquisite sensitivity that adjusts even to the detection of single photons, have evolved for night vision. Cones are less light sensitive but adjust over the range of light intensities found at daylight (more than six orders of magnitude). Moreover, cones are optimized for high temporal stimulus resolution, possessing response times (time to peak ~50 ms) that are about fourfold faster than those of rods (for reviews of photoreceptor physiology, see [1, 2]). Besides day vision, cones underlie other important aspects of vision. In humans, three cone types with different spectral tuning are responsible for color vision, and the retinal spot, built for maximal spatial image resolution, is occupied with cones only. Both rods and cones house the machinery required for photoelectrical transduction in their outer segment, a highly specialized cellular compartment derived from a cilium. The reminder of a stereotypical cilium appears as a narrow, microtubulesupported bridge that separates the outer segment from the main cell body (Fig. 1).

Fig. 1. Schematic of photoreceptor morphology and photoelectrical transduction. Light triggers an enzymatic cascade that leads to the closure of cyclic nucleotide-gated (CNG) channels in the plasma membrane of the photoreceptor outer segment. The CNG channel is associated with an ion exchanger that exports one Ca2+ and one K+ for four imported Na+. In cones, CNG channels are also present in the synaptic terminal. cGMP cyclic 3′,5′-guanosine monophosphate, R rhodopsin, R* light-activated rhodopsin, PDE phosphodiesterase, T transducin.

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Proteins required to maintain structure and function of the outer segment, including CNG channels, are synthesized within the main cell body and, on maturation, are actively transported to their destination. In rods, the outer segment consists of about 1,000 stacked membrane disks that are covered by the plasma membrane. The membrane constituting the disks harbors the proteins involved in photoelectrical transduction, except for the CNG channels. The CNG channels are located in the plasma membrane at high density. In cones, the outer segment is organized differently. Instead of disks, the plasma membrane forms deep invaginations and is the locale of all membrane-bound or integrated proteins involved in photoelectrical transduction. The Activation Phase of the Light Response In the dark, micromolar concentrations of free cGMP keep 1–10% of the CNG channels open in both rods and cones. This open fraction conducts an inward current of about 20 pA that is carried by sodium (Na+) and calcium (Ca2+) ions. This steady inward current (“dark current”) depolarizes photoreceptors to a resting voltage of about −35 mV. In rods, 8–12% of the dark current is carried by Ca2+, while in cones this fraction is 15–25%, twice as high as in rods. The Ca2+ entry is balanced by Ca2+ extrusion through a Na+/K+,Ca2+ exchanger, maintaining a resting Ca2+ concentration of 400–700 nM in both rods and cones. The photoelectrical transduction triggered by light stimulation consists of a complex sequence of biochemical reactions, studied most extensively in rod photoreceptors. On the absorption of a photon, rhodopsin transforms into its active enzymatic form. Active rhodopsin allows the G protein transducin to exchange bound guanosine diphosphate (GDP) for guanosine triphosphate (GTP). GTP-bound transducin activates the effector enzyme phosphodiesterase (PDE), which efficiently hydrolyses cGMP (Fig. 1). The degradation of cGMP and the subsequent closure of CNG channels has two main consequences. First, the photoreceptor hyperpolarizes and releases less of the neurotransmitter glutamate. Second, the interrupted Ca2+ influx into the outer segment leads to a drop in the intracellular Ca2+ concentration due to the continuing export of Ca2+ by the light-insensitive Na+/K+,Ca2+ exchanger. The drop in intracellular Ca2+ is sensed by several Ca2+-binding proteins that accelerate the recovery of the resting state in photoreceptors after light activation and mediate the adaptation of photoreceptors to continuous light exposure (see next section). Photoelectrical transduction in cones is similar to that in rods, and the enzymes involved in light-triggered cGMP degradation display only modest functional differences compared to their rod cousins. Since rods and cones implement functionally similar enzymes in the photoelectrical cascade, which factors are responsible for the differences in light sensitivity, speed, and dynamic response range? It has been suggested that different Ca2+ dynamics in rods and cones partially account for the distinct response properties [2]. Compared with rods, cones display at least a tenfold faster Ca2+ clearance from the outer segment, a lower Ca2+ buffer capacity, and a twofold higher Ca2+ influx through the CNG channels. These differences suggest that the changes in the intracellular Ca2+ concentration are faster and larger in cone outer segments, allowing faster response times and a broader dynamic range of light adaptation.

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Recovery After a Light Stimulus and Adaptation to Continuous Illumination The drop in the intracellular Ca2+ concentration on CNG channel closure is sensed by Ca2+-binding proteins that allow photoreceptors to recover after a light stimulus and to adapt to continuous illumination. Specifically, the protein recoverin is involved in a process that increases the rate of rhodopsin inactivation and guanylate cyclase-activating protein (GCAP) promotes the synthesis of new cGMP. The process of reopening CNG channels and therefore the recovery of the dark current is also Ca2+ dependent. In vitro studies demonstrate that the rod CNG channel is exquisitely sensitive to physiologically relevant levels of Ca2+/calmodulin (e.g., [1, 3]). These data suggest that, in the dark, Ca2+/calmodulin binds to CNG channels and keeps them in a state of low cGMP sensitivity. On the light-triggered drop in intracellular Ca2+, calmodulin dissociates from the CNG channels, thus raising their cGMP sensitivity. As a consequence, less cGMP is required for the channel to reopen. In fact, the cGMP concentration required for half-maximal activation [EC50(cGMP)] of rod channels changes with calmodulin up to twofold from high to low intracellular Ca2+. In contrast, cone CNG channels either respond weakly or are insensitive to modulation by Ca2+/calmodulin. It is possible that another unidentified Ca2+-binding protein modulates the cGMP sensitivity of cone CNG channels over an even broader range [4]. For example, in electropermeabilized cones from the striped bass, the average EC50(cGMP) of CNG channels increases about fourfold from low to high intracellular Ca2+ concentrations. In summary, the Ca2+ permeation and the regulation of activity by intracellular Ca2+ are profoundly different for CNG channels from rods and cones, supporting the notion that the CNG channel is a pivotal determinant of the dynamics of Ca2+ homeostasis in photoreceptors. CNG Channels in the Synaptic Transmission of Cone Photoreceptors In cones, CNG channels are present not only in the outer segment but also in the synaptic terminal, where they might serve an additional function absent in rods. At the photoreceptor dark resting potential of −35 mV, a voltage-activated Ca2+ channel in the synaptic terminal of rods and cones permits continuous Ca2+ entry that sustains a tonic release of glutamate [5]. The Ca2+ channel is characterized by an activation threshold of about −45 mV. When the graded light response reaches −45 mV, the Ca2+ channels close, and in rods, synaptic transmission ceases. In contrast to rods, cones continue synaptic transmission as the light-induced voltage response grows to −70 mV. The discovery of CNG channel clusters in the synaptic terminal of cones offers an explanation for the extended operating range of synaptic transmission [5, 6]. If the clusters are located near release sites, the CNG channels could underlie Ca2+-dependent release of glutamate. In fact, experimental maneuvers that activate CNG channels also trigger exocytotic events and the release of glutamate from the cone terminal. The cGMP required to activate the synaptic CNG channels might be produced by a nitric oxide (NO)-stimulated soluble guanylate cyclase (GC) [7]. Consistent with this model, a NO synthase is present in cone photoreceptors, completing the set of enzymes required for the activation of synaptic CNG channels [7].

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Despite the experimental evidence, a rigorous test of the involvement of CNG channels in synaptic transmission is missing. Such a test will be technically challenging since it requires the selective ablation of synaptic CNG channels. THE MOLECULAR COMPOSITION OF CNG CHANNELS Scientists successfully purified a protein from bovine retina that functionally reconstituted CNG channel activity in artificial liposomes [8]. Partial amino acid sequences derived from the purified protein led to the discovery and functional in vitro expression of the first CNG channel gene in 1989 [9]. The CNG channel protein displays sequence similarity to voltage-gated K+, Na+, and Ca2+ channels, and it was categorized into the same gene superfamily [10]. It contains six transmembrane segments (S1–S6) and a pore-forming stretch of about 20 amino acids (“p region”) that is located between transmembrane segments 5 and 6. The C-terminal region harbors a binding domain for nucleoside 3′,5′-cyclic monophosphates (cNMPs). On heterologous expression, the CNG channel protein displays several of the key properties of the native rod CNG channel but differs in the gating kinetics and the sensitivity to L-cis-diltiazem, a substance that efficiently blocks native channels. The simplest explanation for these functional differences is that the native channel comprises several distinct subunits. In fact, speculation about additional subunits was already stimulated by the presence of a second, larger protein (240 kDa in sodium dodecyl sulfate [SDS]-polyacrylamide gels) in the purified channel preparation (e.g., [11]). Subsequent studies finally identified the 240-kDa protein as a second subunit of rod CNG channels. Coexpression of both channel proteins, later termed CNGA1 (the first subunit) and CNGB1 (the second subunit), recapitulated the properties of native rod CNG channels [12, 13]. CNGB1 exhibits a bipartite structure with a membraneintegrated part that is homologous to CNGA1 (“β′ part”) and a large cytoplasmic N-terminal domain homologous to glutamic acid-rich proteins (GARPs) expressed in rod photoreceptors. The gene encoding the A subunit of cone photoreceptors (CNGA3a) was identified using a cloning strategy that exploited the sequence similarity to the gene encoding CNGA1 [14]. The cone B subunit (CNGB3) was identified by two different approaches. Just before the release of the full sequence of the human genome in 2001, the missing gene was identified in a human expressed sequence tag (EST) database due to its similarity to other CNG channel subunits [15]. At the same time, a genomic region containing CNGB3 was found to be responsible for inherited color blindness among inhabitants of Pingelap, an isolated atoll in the west Pacific [16, 17]. Unlike the counterpart in rod photoreceptors, CNGB3 does not display a bipartite structure and comprises only the membrane-spanning core segment common to all CNG channel subunits. While CNGA1 or CNGA3 form functional channels in heterologous expression systems if expressed alone, CNGB1 or CNGB3 fail to form functional homomeric channels. a

The human genome is comprised of six different CNG channel genes: CNGA1–4, CNGB1, and CNGB3. CNGA2, CNGA4, and CNGB1 encode for the CNG channel in olfactory receptor neurons. CNGB2 has not been assigned.

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Due to this observation, CNGA1 and CNGA3 are often referred to as principal subunits and CNGB1 and CNGB3 as modulatory subunits. How CNGA and CNGB subunits assemble was first understood for the rod CNG channel. The rod CNG channel is a tetramer composed of three CNGA1 subunits and one CNGB1 subunit [18–20]. In contrast, CNG channels in cones adopt a more symmetrical arrangement with two copies of each subunit type, CNGA3 and CNGB3 [21]. It is likely that the four CNG channel subunits arrange such that the p region of each subunit is oriented toward the center of the complex, each contributing to a central membrane-spanning pore. Such an arrangement has been demonstrated for voltagegated K+ channels using x-ray crystallography and is consistent with low-resolution cryoelectron microscopic images of native rod CNG channels [22, 23]. THE BASIC ACTIVATION PROPERTIES OF CNG CHANNELS The CNG channels of rods and cones respond to both cAMP and cGMP but display higher sensitivity to cGMP [13, 24–26] (Fig. 2A). Moreover, cAMP is only a partial agonist for photoreceptor CNG channels, with saturating cAMP concentrations

Fig. 2. Ligand selectivity of rod and cone cyclic nucleotide-gated (CNG) channels. A Heterologously expressed bovine rod (black line) and human cone (gray line) CNG channels are activated by micromolar cyclic 3′,5′-guanosine monophosphate (cGMP) concentrations. In contrast, even millimolar cyclic adenosine monophosphate (cAMP) concentrations activate only a fraction of the maximal cGMP-induced current through both rod and cone CNG channels (data were obtained at a membrane voltage of +80 mV [13, 26]). B Molecular modeling predicts ten interactions between the ligand-binding site of CNG channels and cGMP. C The ligand-binding site of CNG channel subunits is predicted to fold into three α-helices and eight β-strands. Shown is the amino acid sequence of the ligand-binding site in bovine CNGA1. Boxed residues are predicted to interact with cGMP.

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activating only between 10% and 35% of the maximal currents induced by cGMP. Unlike ligand-gated neurotransmitter receptors, CNG channels do not desensitize in the continuous presence of ligand. This feature allows CNG channel activity to faithfully track the cGMP concentration in photoreceptors. Closer inspection of the dose-response relation suggests that CNG channels bind multiple cGMP molecules in a cooperative manner. Specifically, double-logarithmic plots of the activation (log I/Imax vs. log [cGMP]) display a limiting slope of up to about 3.5. This suggests that channel opening requires the binding of 3–4 cGMP molecules [27]. The highly cooperative activation maximizes the sensitivity of CNG channels, and therefore the sensitivity of the photoreceptor dark current, to small changes in the free cGMP concentration. TRANSMEMBRANE TOPOLOGY AND FUNCTIONAL DOMAINS The transmembrane topology of CNG channel A subunits is derived from immunogold labeling and electron microscopy of rod photoreceptors [28, 29], and it is supported by the results of a gene fusion approach using enzyme reporters [30]. According to these studies, both the N- and the C-terminus of A subunits are cytoplasmic, and the segment connecting S5 to the p region is extracellular. Based on sequence similarities, it is assumed that CNG channel A and B subunits adopt a similar transmembrane topology. Several key properties of CNG channels are attributed to specific domains of the channel proteins. These domains deserve a closer look. The Cyclic-Nucleotide-Binding Domain The C-terminal cytoplasmic region of all CNG channel subunits harbors a cNMPbinding site comprised of 80–100 amino acid residues. The three-dimensional structure of the catabolite activator protein (CAP) of Escherichia coli has been used for molecular modeling of the cNMP-binding site in CNG channels (for a detailed discussion, see [31]). The cAMP-binding site of CAP is comprised of three α-helices (A, B, and C) and eight β-strands (β1–β8). The β-strands form a flattened β-roll consisting of two antiparallel β-sheets, each with four strands. The A helix connects to the β-roll, followed by the B and C helix. The ribose and cyclic phosphate interact exclusively with the β-roll, while the purine ring interacts with residues in the β-roll and the C helix. The molecular modeling of the cNMP-binding site in CNG channels predicts ten different interactions between cGMP and the binding pocket of CNGA1 or CNGA3, accomplished by eight different amino acids (Fig. 2B,C). It is thought that interactions with the common ribofuranose moiety are similar for all nucleotides, whereas different interactions with different purine rings control ligand selectivity. In particular, the residues T560 and D604 (in bovine CNGA1) are postulated to be key residues for ligand discrimination. Residue T560, located in β7, is expected to form a hydrogen bond with the amino group of cGMP, probably mediated by a water molecule. In contrast, no interaction is expected to occur between T560 and the purine moiety of cAMP, thus lowering the affinity for cAMP. Consistent with this prediction, the substitution of threonine for alanine dramatically decreases the cGMP sensitivity but has little effect on the cAMP sensitivity. The carboxylate side chain of D604, located in the C helix, presumably shares a single hydrogen bond with either N1 or the amino group in cGMP, while repulsive

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forces are expected with the free electron pair at N1 of cAMP. In agreement with this prediction, the ability of cAMP to activate CNGA1 is enhanced if D604 is replaced with a neutral amino acid (D604Q or D604M), at low pH when D604 is protonated, or when the channel includes CNGB1, which carries a neutral asparagine at the position equivalent to D604. According to the model, T560 and D604 are unlikely to interact simultaneously with the purine ring of cGMP since the two interactions require different conformations of the cyclic nucleotide [32]. It is been speculated that cGMP initially binds in the syn conformation to interact with T560; a switch to the anti conformation then allows the interaction with D604. Moreover, it is also thought that the interaction with D604 initiates a movement of the C helix toward the β-barrel, which is essential for channel gating [33]. The Amino Terminal Domain and Modulation by Calmodulin Deletion studies demonstrated that an unconventional calmodulin-binding site in the N-terminal domain of the β′ part of CNGB1 is essential for the Ca2+/calmodulin sensitivity of rod CNG channels [3, 34]. The detailed mechanism of how binding of Ca2+/calmodulin to this site decreases ligand sensitivity remains unknown. Nevertheless, it appears that disruption of an interaction between CNGB1 and the C-terminal region of CNGA1 is a critical element [34]. Both subunits of cone CNG channels contain conserved calmodulinbinding motifs in their N-termini, and CNGB3 contains an additional calmodulin-binding motif in the C-terminal region (e.g., [35]). Heteromeric cone CNG channels remain sensitive to modulation by Ca2+/calmodulin when either one or the other of the two binding motifs in CNGB3 is deleted but lose sensitivity on deletion of both motifs. The P Region The p region of CNG channels shows high similarity to the p region of voltage-gated K -selective channels, although CNG channels discriminate poorly between monovalent cations [36, 37]. Therefore, not surprisingly, the sequence similarity between the p regions in both channel types ends at the selectivity filter. The crystal structure of KcsA, a bacterial K+-selective channel, provided detailed insight into how selectivity is achieved. In KcsA, the p region consists of an α-helix of about 15 amino acids, followed by a loop of 6 amino acids that forms the selectivity filter in the narrowest part of the pore [22]. It is thought that the gross structure of the pore is conserved in CNG channels, but the structural detail around the selectivity filter is expected to be different. While the selectivity filter in K+ channels comprises only neutral amino acids, CNGA1 and CNGA3 carry one negatively charged glutamate residue in the pore loop. Apparently, pairs of pore glutamates from different subunits interact by sharing a single proton rather than repel each other [38]. Insight into the physiological role of the pore glutamates comes from studies of Ca2+ permeation. Increasing extracellular Ca2+ concentrations progressively impedes the current of monovalent ions through CNG channels (Fig. 3A). At physiological Ca2+ concentrations, CNG channel currents are reduced to a few percent of the maximal value found under Ca2+-free conditions. For example, the single-channel conductance of rod CNG channels drops from about 25 pS in a Ca2+-free saline to less than 1 pS in the presence of 1 mM extracellular Ca2+ [39, 40]. +

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Fig. 3. Blockage of cyclic nucleotide-gated (CNG) channels by extracellular calcium. A Increasing the extracellular Ca2+ concentration successively impedes currents through homomeric CNGA1 (black dashed line) and CNGA3 (gray dashed line) channels as well as currents through heteromeric rod (CNGA1/CNGB1; black solid line) and cone (CNGA3/CNGB3; gray solid line) channels. Data were obtained on heterologous expression of CNG channel subunits at a membrane voltage of −60 mV [13, 26]. B Each subunit in rod (left) and cone (right) CNG channels contains an α-helix followed by a loop in the pore-forming region. The pore loop in CNG channel A subunits contains a glutamate that participates in the formation of the ion-binding site within the pore (E). B subunits carry a glycine (G) instead of glutamate in the pore loop.

The effect of Ca2+, and the similar effect of Mg2+, on CNG channel currents can be explained by a model in which permeating monovalent and divalent ions compete for a common cation-binding site within the channel pore. Since divalent ions are bound more strongly by the binding site, they occlude the permeation pathway for monovalent ions. The fact the Ca2+ blockage can be described by a simple binding isotherm demonstrates the presence of a single Ca2+-binding site within the pore [41]. The Ca2+-binding site is apparently formed by the pore glutamates of CNGA1 and CNGA3 since the replacement of glutamate by the neutral residue glutamine dramatically reduces the Ca2+ blockage [42]. Homomeric channels composed only of CNGA1 generally display a higher sensitivity for Ca2+ blockage than those composed only of CNGA3. An extensive mutagenesis study showed that all aspects of Ca2+ blockage can be transferred from one channel to another by swapping the p region and the two adjacent transmembrane segments (S5–p region–S6) of the channel-forming subunits. Therefore, the S5–p region–S6 segment is considered the basic pore module that governs both blockage and ion permeation in CNG channels [41]. CNGB1 and CNGB3 do not carry a negatively charged pore loop glutamate but instead have a neutral glycine. Thus, native rod channels have a total of three pore glutamates, and native cone channels presumably a total of two pore glutamates (Fig. 3B). The fact that the pore affinity for Ca2+ is reduced in channels with incorporated B subunits is likely to be due to the reduction in the pore’s charge density [31]. The high affinity of the CNG channel pore for Ca2+ has two important consequences for the physiology of photoreceptors. First, it ensures that the pore of CNG channels is preferentially occupied with Ca2+ rather than monovalent ions. This allows a relative Ca2+ flux of 10–25%, even though Ca2+ constitutes less than 2% of the extracellular cations. Second, Ca2+ blockage

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reduces the contribution of each single channel to the total dark current, making it less sensitive to single-channel current fluctuations and thereby increasing the signal-tonoise ratio of photoreceptor currents. This aspect is of special importance for rod photoreceptors since they act as high-sensitivity photon detectors. Interestingly, amino acid substitutions in or near the pore glutamate not only change the permeation properties but also the gating behavior of CNG channels (e.g., [26]). This observation indicates that conformational rearrangements of the pore around the selectivity filter are also involved in channel gating. The GARP Domain of CNGB1 CNGB1 is unique among CNG channel subunits because it contains a large N-terminal cytoplasmic domain that is almost identical to two soluble rod-specific GARPs. The GARPs are alternatively spliced forms of a single gene different from that encoding CNGB1. The most conserved structural elements in GARPs, four short proline-rich repeats of about 15 amino acids in their N-terminus, were used as bait to test for interactions of CNGB1 with other rod proteins using peptide affinity chromatography [43]. This study found that the proline-rich repeats from GARPs interact with PDE, GC, and the retina-specific ATP-binding cassette receptor (ABCR) transporter. The ABCR transporter is also known as the rim protein as it is distributed along the rim region of rod discs. The interaction between GARPs and ABCR and GC has been called into question by the results of a second approach to identify GARP-interacting proteins [44]. In immunoprecipitation experiments, neither ABCR nor GC was pulled down with GARP-specific antibodies, even when GARP-binding partners were covalently attached using cross-linking reagents. Instead, peripherin, another protein located at the disk rim, was shown to interact with the shorter GARP splice variant and the GARP domain of CNGB1 (Fig. 4). The hydrodynamic properties of GARPs and little secondary and tertiary structure are consistent with an elongated, unfolded GARP domain of CNGB1 that is able to span the 10-nm gap between the rod plasma membrane and the disk rim [45]. The interaction between CNGB1 and proteins at the disk rim would align the CNG channels in stacked circles along the outer segment. Indeed, a nonuniform distribution of CNG channels in the outer segment of rod photoreceptors has been reported [46]. This suggests that the interaction between CNGB1 and disk rim proteins supports the flat appearance and the arrangement of membrane disks in the outer segment; however, mice lacking CNGB1 form rods of normal morphology [47]. Due to the high density of glutamate residues in the GARP domain of CNGB1 (137 of 571 residues in bovine CNGB1), this domain functions as a low-affinity Ca2+ buffer. It has been speculated that the GARP domain of CNGB1 guides Ca2+ ions from the intracellular CNG channel pore to the disk surface, the locale of the Ca2+-binding proteins that regulate the sensitivity and the kinetics of the photoresponse [45]. CNG CHANNELS ARE COMPONENTS OF LARGER PROTEIN COMPLEXES The interaction of peripherin and the GARP domain of CNGB1 argue against a model in which CNG channels diffuse freely and are isolated in the photoreceptor membrane. In fact, further biochemical and pharmacological data suggest that CNG channels of

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Fig. 4. Rod cyclic nucleotide-gated (CNG) channels interact with peripherin, a Na+/K+,Ca2+ exchanger, and a protein tyrosine kinase (PTK). See text for details. ABCR ATP-binding cassette.

photoreceptors are components of even larger protein complexes. In both rods and cones, CNG channels are intimately associated with the Na+/K+,Ca2+ exchanger via their A subunits [44, 48]. The juxtaposition of the channel and the exchanger suggests that Ca2+ dynamics inside the cell are localized to microdomains in the vicinity of the channel (Fig. 4). An unexpected finding suggests that CNG channels in photoreceptors are also associated with a protein tyrosine kinase (PTK). Genistein, an inhibitor of PTKs, reversibly slows the gating kinetics and reduces the maximal current responses of native CNG channels in the absence of ATP (i.e., independent of phosphorylation) [49]. The effect of genistein on CNG channels is ameliorated by other PTK inhibitors that do not otherwise affect CNG channels. In conclusion of this finding, it was suggested that genistein binds to the PTK rather than the CNG channel. It is thought that PTK undergoes conformational changes on binding of genistein that are transferred to the tightly associated CNG channel, thereby modifying channel function. At this time, however, a physical association between PTKs and CNG channels has not been demonstrated. MODULATION BY PHOSPHORYLATION AND ALL-TRANS RETINAL The cGMP sensitivity of photoreceptor CNG channels is modulated by tyrosine and serine/threonine phosphorylation. The effects of tyrosine phosphorylation on rod CNG channels have been extensively studied in vitro [24, 49]. Heteromeric CNGA1/CNGB1 channels expressed in Xenopus oocytes show a roughly twofold increase in cGMP sensitivity developing over time after patch excision. This increase was attributed to the spontaneous dephosphorylation of single tyrosine residues in the cNMP-binding site of both CNGA1 and CNGB1 by protein tyrosine phosphatases (PTPs). A signaling pathway that may regulate tyrosine phosphorylation of rod CNG channels in vivo involves insulin-like growth factor 1 (IGF-1) [49]. IGF-1 is released from the pigment epithelium and increases the cGMP sensitivity of CNG channels in rod photoreceptors. IGF-1 is thought to exert the effect on CNG channels through the regulation of a PTP.

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Besides tyrosine phosphorylation, also serine/threonine phosphorylation has been implicated in the regulation of the ligand sensitivity of retinal CNG channels. Repeated dose-response measurements with CNG channels in excised patches from rod outer segments revealed a slow increase in cGMP sensitivity over time, characterized by an up to tenfold decrease in the EC50 (cGMP) [50]. The enhancement of ligand sensitivity was slowed by ATP or inhibitors of serine/threonine phosphatases and accelerated by purified type I phosphatase. Similarly, a threefold decrease in cGMP sensitivity was observed for heterologously expressed CNGA3 channels on exposure to phorbol esters, substances that stimulate protein kinase C (PKC) [51]. This effect is due to the phosphorylation of two serine residues, also located in the cNMP-binding site. The cGMP sensitivity of cone CNG channels, at least in chicken, is under the control of a circadian rhythm [52]. During the subjective night, the sensitivity is approximately twofold higher than during the subjective day. This circadian modulation is apparently driven by rhythms in the activities of the extracellular receptor kinase (ERK) form of mitogen-activated protein kinase and the Ca2+/calmodulin-dependent protein kinase II (CaMKII), but the detailed mechanism underlying this control is unknown. Besides phosphorylation, rod CNG channels are also inhibited by the phospholipid Phosphoinositol diphosphate (PIP2) and all-trans retinal, the photoisomerized pigment released from rhodopsin after light activation [53]. All-trans retinal directly inhibits the homomeric CNGA1 and heteromeric CNGA1/CNGB1 channels at nanomolar concentrations, probably due to a decrease in the open probability. The effects observed in a heterologous expression system might also apply to native rod CNG channels, when bright illumination is expected to lead to elevated concentrations of photoisomerized pigment in the outer segment. SYNTHESIS, MATURATION, AND TARGETING OF CNG CHANNELS The predominant form of CNGA1 in bovine rod outer segments has an apparent molecular mass of 63 kDa, which is significantly less then the molecular mass predicted from the full-length complementary DNA or the apparent molecular mass of heterologously expressed CNGA1 (78 kDa). This discrepancy in size is due to a photoreceptor-specific proteolytic process by which the 92 N-terminal amino acids of CNGA1 are removed [28]. Similar processing of the rod A subunits seems to be common across species, and at least the chicken cone A subunit also undergoes posttranslational cleavage [14, 54]. Another posttranslational modification of CNGA1 and CNGA3 is the glycosylation of an asparagine residue in the extracellular loop between S5 and the p region [29, 55]. In contrast to the CNG channel A subunits, CNGB1 and CNGB3 are unglycosylated. Currently, the functions of proteolytic cleavage and glycosylation are unknown since neither affects channel activity. The posttranslational modification might be required for channel targeting or for the interaction of CNG channels with other photoreceptor proteins. For proper targeting, CNG channels need to be translocated from their site of synthesis in the membrane of the endoplasmic reticulum (ER) within the main cell body to the plasma membrane of the outer segment. Impaired plasma membrane targeting in heterologous expression systems has been reported for a variety of mutant CNGA1 and CNGA3 subunits found in patients with either rod or cone dysfunction, respectively. Most of these mutants fold improperly and thus fail to leave the ER [55–57].

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In a naturally occurring CNGA1 mutant with an amino acid substitution and truncation C-terminal to the cNMP-binding site (R654D-stop in human CNGA1), a more specific mechanism of ER retention seems to apply. Two studies investigated the mechanism of ER retention for this mutant, but with conflicting results. One study found that the mutation generates an ER retention signal itself, while another study provided evidence that the mutated CNGA1 is unable to mask an ER retention signal in CNGB1 [58, 59]. At present it is unclear how the conflicting results can be reconciled. A glimpse of how CNG channels are transported into the outer segment was obtained in a study using subunits of the olfactory CNG channels (CNGA2 and CNGB1b, a splice variant of CNGB1 that lacks the GARP domain) expressed in a ciliated cell line [60]. In this study, the ciliary transport of CNGA2 is dependent on CNGB1b and, more specifically, on a short sequence of four amino acids within this subunit (RVxP, with x representing a variable amino acid). A similar mechanism might apply to rod CNG channels and could explain why rods from mice lacking CNGB1 have decreased levels of CNGA1 in the outer segment [47]. VISUAL DYSFUNCTION CAUSED BY MUTANT CNG CHANNEL GENES Mutations in the genes for CNG channels of photoreceptors are associated with the hereditary visual diseases retinitis pigmentosa (RP), achromatopsia, and cone dystrophy [16, 17, 57, 61–68] (see also Appendix). Retinitis pigmentosa is a clinically and genetically heterogeneous group of diseases characterized by night blindness, a progressive loss of the peripheral visual field, and eventual loss of central vision, resulting in blindness. These symptoms reflect early dysfunction and degeneration of rod photoreceptors, followed by a slower degeneration of cone photoreceptors that proceeds from the periphery to the center of the visual field. Mutations in the CNGA1 or CNGB1 gene only account for a few percent of autosomal recessive RP. Achromatopsia is a recessive, nonprogressive disease resulting from the dysfunction of cone photoreceptors. Symptoms include absence of color vision, light sensitivity (photophobia), and poor visual acuity. Of all cases of achromatopsia, 20–30% are caused by CNGA3 mutations and 40–50% by mutations in CNGB3 [69]. Moreover, in some instances, mutations in CNGA3 or CNGB3 result in cone dystrophy, a disease related to achromatopsia but characterized by the progressive loss of cone function and sometimes the progressive loss of rod function. The vast number of deleterious single amino acid substitutions in CNGA3 (Fig. 5) indicates that there is little tolerance for sequence variations in CNG channel A subunits with respect to photoreceptor function. Mutations do not, however, necessarily lead to nonfunctional CNG channels. Some mutant channel proteins, in particular those exclusively found in patients with residual photoreceptor function (Fig. 5, underlined mutations), are expected to form channels, although with functional alterations. A mild form of achromatopsia with considerably preserved cone function is caused by the heterozygous mutations T224R and T369S in CNGA3 (A3T224R and A3T369S) [26]. The patients are able to discriminate saturated but not desaturated colors. Psychophysical and electroretinographical analyses showed that the cone system is characterized by lower light sensitivity and perturbed signal transfer from cones to postsynaptic neurons. Patch-clamp analysis of heterologously expressed subunits revealed that only A3T369S

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Fig. 5. Mutations in cone photoreceptor B subunit (CNGA3) are associated with cone dysfunction. See text for details. Underlined amino acid substitutions are found in patients with residual cone function. cNMP nucleoside 3′,5′-cyclic monophosphate, Fs frame shift mutation, del deletion.

produces functional channels, although with grossly altered permeation of monovalent and divalent ions, gating, and ligand sensitivity. Coexpression of wild-type CNGB3 with A3T369S restored most of the native properties, except for the altered Ca2+ permeation. The properties of A3T369S/CNGB3 channels suggest that the mild form of achromatopsia results from relatively subtle changes in ion flux through the cone CNG channel. Another channel defect appears to be common in patients with cone dysfunction. Several missense mutations associated with cone dystrophy (A3N471S, A3R563H, B3R403Q) or complete achromatopsia (B3F525N, B3D633G, and the Pingelap mutation B3S435F) have been reported to produce channels with moderately increased cGMP sensitivity [25, 66, 70, 71]. These findings are surprising given the severe cone defects in the patients but might reflect the fact that the functional integrity of photoreceptors relies on a precisely tuned transduction machinery. How a similar functional defect in different mutant channels can produce different clinical phenotypes remains unresolved. Intriguingly, the absence or malfunction of CNG channels often leads to photoreceptor loss and even retinal degeneration. Our understanding of the trigger, the detailed mechanisms underlying retinal degeneration, and in particular the role of CNG channels in these processes is incomplete. We have to assume that the survival of photoreceptors critically depends on the structural and functional integrity of their enzymatic machinery. The discovery of efficient CNG channel blockers (e.g., [72]) opens an avenue for the treatment of retinal diseases caused by abnormally high CNG channel activity.

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REFERENCES 1. Lamb, T. D., Pugh, E. N., Jr. (2000). Phototransduction in vertebrate rods and cones: molecular mechanisms of amplification, recovery and light adaptation. In: Handbook of biological physics (Stavenga, D. G., de Grip, W. J., Pugh, E. N., Jr., eds.), Vol. 3, pp. 183–254. Elsevier, Amsterdam. 2. Miller, J. L., Korenbrot, J. I. (1994). Differences in calcium homeostasis between retinal rod and cone photoreceptors revealed by the effects of voltage on the cGMP-gated conductance in intact cells. J Gen Physiol 104, 909–940. 3. Weitz, D., Zoche, M., Muller, F., Beyermann, M., Korschen, H. G., Kaupp, U. B., Koch, K. W. (1998). Calmodulin controls the rod photoreceptor CNG channel through an unconventional binding site in the N-terminus of the beta-subunit. EMBO J 17, 2273–2284. 4. Rebrik, T. I., Korenbrot, J. I. (1998). In intact cone photoreceptors, a Ca2+-dependent, diffusible factor modulates the cGMP-gated ion channels differently than in rods. J Gen Physiol 112, 537–548. 5. Rieke, F., Schwartz, E. A. (1994). A cGMP-gated current can control exocytosis at cone synapses. Neuron 13, 863–873. 6. Savchenko, A., Barnes, S., Kramer, R. H. (1997). Cyclic-nucleotide-gated channels mediate synaptic feedback by nitric oxide. Nature 390, 694–698. 7. Koch, K. W., Lambrecht, H. G., Haberecht, M., Redburn, D., Schmidt, H. H. (1994). Functional coupling of a Ca2+/calmodulin-dependent nitric oxide synthase and a soluble guanylyl cyclase in vertebrate photoreceptor cells. EMBO J 13, 3312–3320. 8. Cook, N. J., Hanke, W., Kaupp, U. B. (1987). Identification, purification, and functional reconstitution of the cyclic GMP-dependent channel from rod photoreceptors. Proc Natl Acad Sci U S A 84, 585–589. 9. Kaupp, U. B., Niidome, T., Tanabe, T., Terada, S., Bonigk, W., Stuhmer, W., Cook, N. J., Kangawa, K., Matsuo, H., Hirose, T., et al. (1989). Primary structure and functional expression from complementary DNA of the rod photoreceptor cyclic GMP-gated channel. Nature 342, 762–766. 10. Jan, L. Y., Jan, Y. N. (1990). A superfamily of ion channels. Nature 345, 672. 11. Molday, L. L., Cook, N. J., Kaupp, U. B., Molday, R. S. (1990). The cGMP-gated cation channel of bovine rod photoreceptor cells is associated with a 240-kDa protein exhibiting immunochemical cross-reactivity with spectrin. J Biol Chem 265, 18690–18695. 12. Chen, T. Y., Peng, Y. W., Dhallan, R. S., Ahamed, B., Reed, R. R., Yau, K. W. (1993). A new subunit of the cyclic nucleotide-gated cation channel in retinal rods. Nature 362, 764–767. 13. Korschen, H. G., Illing, M., Seifert, R., Sesti, F., Williams, A., Gotzes, S., Colville, C., Muller, F., Dose, A., Godde, M., et al. (1995). A 240 kDa protein represents the complete beta subunit of the cyclic nucleotide-gated channel from rod photoreceptor. Neuron 15, 627–636. 14. Bonigk, W., Altenhofen, W., Muller, F., Dose, A., Illing, M., Molday, R. S., Kaupp, U. B. (1993). Rod and cone photoreceptor cells express distinct genes for cGMP-gated channels. Neuron 10, 865–877. 15. Gerstner, A., Zong, X., Hofmann, F., Biel, M. (2000). Molecular cloning and functional characterization of a new modulatory cyclic nucleotide-gated channel subunit from mouse retina. J Neurosci 20, 1324–1332. 16. Sundin, O. H., Yang, J. M., Li, Y., Zhu, D., Hurd, J. N., Mitchell, T. N., Silva, E. D., Maumenee, I. H. (2000). Genetic basis of total colourblindness among the Pingelapese islanders. Nat Genet 25, 289–293. 17. Kohl, S., Baumann, B., Broghammer, M., Jagle, H., Sieving, P., Kellner, U., Spegal, R., Anastasi, M., Zrenner, E., Sharpe, L. T., Wissinger, B. (2000). Mutations in the CNGB3 gene

240

18. 19. 20. 21. 22.

23.

24.

25.

26.

27.

28.

29.

30.

31. 32. 33. 34. 35.

Tränkner encoding the beta-subunit of the cone photoreceptor cGMP-gated channel are responsible for achromatopsia (ACHM3) linked to chromosome 8q21. Hum Mol Genet 9, 2107–2116. Weitz, D., Ficek, N., Kremmer, E., Bauer, P. J., Kaupp, U. B. (2002). Subunit stoichiometry of the CNG channel of rod photoreceptors. Neuron 36, 881–889. Zheng, J., Trudeau, M. C., Zagotta, W. N. (2002). Rod cyclic nucleotide-gated channels have a stoichiometry of three CNGA1 subunits and one CNGB1 subunit. Neuron 36, 891–896. Zhong, H., Molday, L. L., Molday, R. S., Yau, K. W. (2002). The heteromeric cyclic nucleotidegated channel adopts a 3A:1B stoichiometry. Nature 420, 193–198. Peng, C., Rich, E. D., Varnum, M. D. (2004). Subunit configuration of heteromeric cone cyclic nucleotide-gated channels. Neuron 42, 401–410. Doyle, D. A., Morais Cabral, J., Pfuetzner, R. A., Kuo, A., Gulbis, J. M., Cohen, S. L., Chait, B. T., MacKinnon, R. (1998). The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science 280, 69–77. Higgins, M. K., Weitz, D., Warne, T., Schertler, G. F., Kaupp, U. B. (2002). Molecular architecture of a retinal cGMP-gated channel: the arrangement of the cytoplasmic domains. EMBO J 21, 2087–2094. Molokanova, E., Krajewski, J. L., Satpaev, D., Luetje, C. W., Kramer, R. H. (2003). Subunit contributions to phosphorylation-dependent modulation of bovine rod cyclic nucleotide-gated channels. J Physiol 552, 345–356. Peng, C., Rich, E. D., Varnum, M. D. (2003). Achromatopsia-associated mutation in the human cone photoreceptor cyclic nucleotide-gated channel CNGB3 subunit alters the ligand sensitivity and pore properties of heteromeric channels. J Biol Chem 278, 34533–34540. Trankner, D., Jagle, H., Kohl, S., Apfelstedt-Sylla, E., Sharpe, L. T., Kaupp, U. B., Zrenner, E., Seifert, R., Wissinger, B. (2004). Molecular basis of an inherited form of incomplete achromatopsia. J Neurosci 24, 138–147. Ruiz, M., Brown, R. L., He, Y., Haley, T. L., Karpen, J. W. (1999). The single-channel doseresponse relation is consistently steep for rod cyclic nucleotide-gated channels: implications for the interpretation of macroscopic dose-response relations. Biochemistry 38, 10642– 10648. Molday, R. S., Molday, L. L., Dose, A., Clark-Lewis, I., Illing, M., Cook, N. J., Eismann, E., Kaupp, U. B. (1991). The cGMP-gated channel of the rod photoreceptor cell characterization and orientation of the amino terminus. J Biol Chem 266, 21917–21922. Wohlfart, P., Haase, W., Molday, R. S., Cook, N. J. (1992). Antibodies against synthetic peptides used to determine the topology and site of glycosylation of the cGMP-gated channel from bovine rod photoreceptors. J Biol Chem 267, 644–648. Henn, D. K., Baumann, A., Kaupp, U. B. (1995). Probing the transmembrane topology of cyclic nucleotide-gated ion channels with a gene fusion approach. Proc Natl Acad Sci U S A 92, 7425–7429. Kaupp, U. B., Seifert, R. (2002). Cyclic nucleotide-gated ion channels. Physiol Rev 82, 769–824. Varnum, M. D., Black, K. D., Zagotta, W. N. (1995). Molecular mechanism for ligand discrimination of cyclic nucleotide-gated channels. Neuron 15, 619–25. Mazzolini, M., Punta, M., Torre, V. (2002). Movement of the C-helix during the gating of cyclic nucleotide-gated channels. Biophys J 83, 3283–3295. Trudeau, M. C., Zagotta, W. N. (2002). Mechanism of calcium/calmodulin inhibition of rod cyclic nucleotide-gated channels. Proc Natl Acad Sci U S A 99, 8424–8429. Peng, C., Rich, E. D., Thor, C. A., Varnum, M. D. (2003). Functionally important calmodulinbinding sites in both NH2- and COOH-terminal regions of the cone photoreceptor cyclic nucleotide-gated channel CNGB3 subunit. J Biol Chem 278, 24617–24623.

Transduction Channels of Rod and Cone Photoreceptors

241

36. Menini, A. (1990). Currents carried by monovalent cations through cyclic GMP-activated channels in excised patches from salamander rods. J Physiol 424, 167–185. 37. Picones, A., Korenbrot, J. I. (1992). Permeation and interaction of monovalent cations with the cGMP-gated channel of cone photoreceptors. J Gen Physiol 100, 647–673. 38. Morrill, J. A., MacKinnon, R. (1999). Isolation of a single carboxyl-carboxylate proton binding site in the pore of a cyclic nucleotide-gated channel. J Gen Physiol 114, 71–83. 39. Bodoia, R. D., Detwiler, P. B. (1985). Patch-clamp recordings of the light-sensitive dark noise in retinal rods from the lizard and frog. J Physiol 367, 183–216. 40. Haynes, L. W., Kay, A. R., Yau, K. W. (1986). Single cyclic GMP-activated channel activity in excised patches of rod outer segment membrane. Nature 321, 66–70. 41. Seifert, R., Eismann, E., Ludwig, J., Baumann, A., Kaupp, U. B. (1999). Molecular determinants of a Ca2+-binding site in the pore of cyclic nucleotide-gated channels: S5/S6 segments control affinity of intrapore glutamates. EMBO J 18, 119–130. 42. Eismann, E., Muller, F., Heinemann, S. H., Kaupp, U. B. (1994). A single negative charge within the pore region of a cGMP-gated channel controls rectification, Ca2+ blockage, and ionic selectivity. Proc Natl Acad Sci U S A 91, 1109–1113. 43. Korschen, H. G., Beyermann, M., Muller, F., Heck, M., Vantler, M., Koch, K. W., Kellner, R., Wolfrum, U., Bode, C., Hofmann, K. P., Kaupp, U. B. (1999). Interaction of glutamicacid-rich proteins with the cGMP signalling pathway in rod photoreceptors. Nature 400, 761–766. 44. Poetsch, A., Molday, L. L., Molday, R. S. (2001). The cGMP-gated channel and related glutamic acid-rich proteins interact with peripherin-2 at the rim region of rod photoreceptor disc membranes. J Biol Chem 276, 48009–40016. 45. Batra-Safferling, R., Abarca-Heidemann, K., Korschen, H. G., Tziatzios, C., Stoldt, M., Budyak, I., Willbold, D., Schwalbe, H., Klein-Seetharaman, J., Kaupp, U. B. (2006). Glutamic acid-rich proteins of rod photoreceptors are natively unfolded. J Biol Chem 281, 1449– 1460. 46. Karpen, J. W., Loney, D. A., Baylor, D. A. (1992). Cyclic GMP-activated channels of salamander retinal rods: spatial distribution and variation of responsiveness. J Physiol 448, 257–274. 47. Huttl, S., Michalakis, S., Seeliger, M., Luo, D. G., Acar, N., Geiger, H., Hudl, K., Mader, R., Haverkamp, S., Moser, M., Pfeifer, A., Gerstner, A., Yau, K. W., Biel, M. (2005). Impaired channel targeting and retinal degeneration in mice lacking the cyclic nucleotide-gated channel subunit CNGB1. J Neurosci 25, 130–138. 48. Kang, K., Bauer, P. J., Kinjo, T. G., Szerencsei, R. T., Bonigk, W., Winkfein, R. J., Schnetkamp, P. P. (2003). Assembly of retinal rod or cone Na(+)/Ca(2+)-K(+) exchanger oligomers with cGMP-gated channel subunits as probed with heterologously expressed cDNAs. Biochemistry 42, 4593–4600. 49. Molokanova, E., Kramer, R. H. (2001). Mechanism of inhibition of cyclic nucleotide-gated channel by protein tyrosine kinase probed with genistein. J Gen Physiol 117, 219–234. 50. Gordon, S. E., Brautigan, D. L., Zimmerman, A. L. (1992). Protein phosphatases modulate the apparent agonist affinity of the light-regulated ion channel in retinal rods. Neuron 9, 739–748. 51. Muller, F., Vantler, M., Weitz, D., Eismann, E., Zoche, M., Koch, K. W., Kaupp, U. B. (2001). Ligand sensitivity of the 2 subunit from the bovine cone cGMP-gated channel is modulated by protein kinase C but not by calmodulin. J Physiol 532, 399–409. 52. Ko, G. Y., Ko, M. L., Dryer, S. E. (2001). Circadian regulation of cGMP-gated cationic channels of chick retinal cones. Erk MAP Kinase and Ca2+/calmodulin-dependent protein kinase II. Neuron 29, 255–266.

242

Tränkner

53. McCabe, S. L., Pelosi, D. M., Tetreault, M., Miri, A., Nguitragool, W., Kovithvathanaphong, P., Mahajan, R., Zimmerman, A. L. (2004). All-trans-retinal is a closed-state inhibitor of rod cyclic nucleotide-gated ion channels. J Gen Physiol 123, 521–531. 54. Molday, R. S., Reid, D. M., Connell, G., Molday, L. L. (1992). Molecular properties of the cGMP-gated cation channel of rod photoreceptor cells as probed with monoclonal antibodies. In: Signal transduction in photoreceptor cells (Hargrave, P. A., Hofmann, K. P., Kaupp, U. B., eds.), pp. 180. Springer-Verlag, Berlin. 55. Faillace, M. P., Bernabeu, R. O., Korenbrot, J. I. (2004). Cellular processing of cone photoreceptor cyclic GMP-gated ion channels: a role for the S4 structural motif. J Biol Chem 279, 22643–22653. 56. Patel, K. A., Bartoli, K. M., Fandino, R. A., Ngatchou, A. N., Woch, G., Carey, J., Tanaka, J. C. (2005). Transmembrane S1 mutations in CNGA3 from achromatopsia 2 patients cause loss of function and impaired cellular trafficking of the cone CNG channel. Invest Ophthalmol Vis Sci 46, 2282–2290. 57. Dryja, T. P., Finn, J. T., Peng, Y. W., McGee, T. L., Berson, E. L., Yau, K. W. (1995). Mutations in the gene encoding the alpha subunit of the rod cGMP-gated channel in autosomal recessive retinitis pigmentosa. Proc Natl Acad Sci U S A 92, 10177–10181. 58. Mallouk, N., Ildefonse, M., Pages, F., Ragno, M., Bennett, N. (2002). Basis for intracellular retention of a human mutant of the retinal rod channel alpha subunit. J Membr Biol 185, 129–136. 59. Trudeau, M. C., Zagotta, W. N. (2002). An intersubunit interaction regulates trafficking of rod cyclic nucleotide-gated channels and is disrupted in an inherited form of blindness. Neuron 34, 197–207. 60. Jenkins, P. M., Hurd, T. W., Zhang, L., McEwen, D. P., Brown, R. L., Margolis, B., Verhey, K. J., Martens, J. R. (2006). Ciliary targeting of olfactory CNG channels requires the CNGB1b subunit and the kinesin-2 motor protein, KIF17. Curr Biol 16, 1211–1216. 61. Paloma, E., Martinez-Mir, A., Garcia-Sandoval, B., Ayuso, C., Vilageliu, L., GonzalezDuarte, R., Balcells, S. (2002). Novel homozygous mutation in the alpha subunit of the rod cGMP gated channel (CNGA1) in two Spanish sibs affected with autosomal recessive retinitis pigmentosa. J Med Genet 39, E66. 62. Bareil, C., Hamel, C. P., Delague, V., Arnaud, B., Demaille, J., Claustres, M. (2001). Segregation of a mutation in CNGB1 encoding the beta-subunit of the rod cGMP-gated channel in a family with autosomal recessive retinitis pigmentosa. Hum Genet 108, 328–334. 63. Wissinger, B., Gamer, D., Jagle, H., Giorda, R., Marx, T., Mayer, S., Tippmann, S., Broghammer, M., Jurklies, B., Rosenberg, T., Jacobson, S. G., Sener, E. C., Tatlipinar, S., Hoyng, C. B., Castellan, C., Bitoun, P., Andreasson, S., Rudolph, G., Kellner, U., Lorenz, B., Wolff, G., Verellen-Dumoulin, C., Schwartz, M., Cremers, F. P., Apfelstedt-Sylla, E., Zrenner, E., Salati, R., Sharpe, L. T., Kohl, S. (2001). CNGA3 mutations in hereditary cone photoreceptor disorders. Am J Hum Genet 69, 722–737. 64. Johnson, S., Michaelides, M., Aligianis, I. A., Ainsworth, J. R., Mollon, J. D., Maher, E. R., Moore, A. T., Hunt, D. M. (2004). Achromatopsia caused by novel mutations in both CNGA3 and CNGB3. J Med Genet 41, e20. 65. Rojas, C. V., Maria, L. S., Santos, J. L., Cortes, F., Alliende, M. A. (2002). A frameshift insertion in the cone cyclic nucleotide gated cation channel causes complete achromatopsia in a consanguineous family from a rural isolate. Eur J Hum Genet 10, 638–642. 66. Okada, A., Ueyama, H., Toyoda, F., Oda, S., Ding, W. G., Tanabe, S., Yamade, S., Matsuura, H., Ohkubo, I., Kani, K. (2004). Functional role of hCngb3 in regulation of human cone cng channel: effect of rod monochromacy-associated mutations in hCNGB3 on channel function. Invest Ophthalmol Vis Sci 45, 2324–2332.

Transduction Channels of Rod and Cone Photoreceptors

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67. Nishiguchi, K. M., Sandberg, M. A., Gorji, N., Berson, E. L., Dryja, T. P. (2005). Cone cGMP-gated channel mutations and clinical findings in patients with achromatopsia, macular degeneration, and other hereditary cone diseases. Hum Mutat 25, 248–258. 68. Goto-Omoto, S., Hayashi, T., Gekka, T., Kubo, A., Takeuchi, T., Kitahara, K. (2006). Compound heterozygous CNGA3 mutations (R436W, L633P) in a Japanese patient with congenital achromatopsia. Vis Neurosci 23, 395–402. 69. Kohl, S., Varsanyi, B., Antunes, G. A., Baumann, B., Hoyng, C. B., Jagle, H., Rosenberg, T., Kellner, U., Lorenz, B., Salati, R., Jurklies, B., Farkas, A., Andreasson, S., Weleber, R. G., Jacobson, S. G., Rudolph, G., Castellan, C., Dollfus, H., Legius, E., Anastasi, M., Bitoun, P., Lev, D., Sieving, P. A., Munier, F. L., Zrenner, E., Sharpe, L. T., Cremers, F. P., Wissinger, B. (2005). CNGB3 mutations account for 50% of all cases with autosomal recessive achromatopsia. Eur J Hum Genet 13, 302–308. 70. Bright, S. R., Brown, T. E., Varnum, M. D. (2005). Disease-associated mutations in CNGB3 produce gain of function alterations in cone cyclic nucleotide-gated channels. Mol Vis 11, 1141–1150. 71. Liu, C., Varnum, M. D. (2005). Functional consequences of progressive cone dystrophyassociated mutations in the human cone photoreceptor cyclic nucleotide-gated channel CNGA3 subunit. Am J Physiol Cell Physiol 289, C187–C198. 72. Brown, R. L., Haley, T. L., West, K. A., Crabb, J. W. (1999). Pseudechetoxin: a peptide blocker of cyclic nucleotide-gated ion channels. Proc Natl Acad Sci U S A 96, 754–759.

APPENDIX Visual Dysfunction Caused by Mutant CNG Channel Genes Mutations in cyclic nucleotide-gated (CNG) channel genes can cause the malfunction and degeneration of photoreceptors and, concomitantly, partial or complete blindness. Mutations in the genes encoding the CNG channel subunits of rod photoreceptors— CNGA1 and CNGB1—account for a few percent of recessive retinitis pigmentosa (RP) [1–3]. RP is characterized by night blindness, a progressive loss of the peripheral visual field, and eventual loss of central vision, resulting in blindness. These symptoms reflect early dysfunction and degeneration of rod photoreceptors, followed by a slower degeneration of cone photoreceptors that proceeds from the periphery to the center of the visual field (for review, see [4, 5]). Mutations in the genes for the CNG channel subunits of cones— CNGA3 and CNGB3—can cause achromatopsia. Achromatopsia is a recessive, nonprogressive disease resulting from the dysfunction of cone photoreceptors. Symptoms include absence of color vision, light sensitivity (photophobia), and poor visual acuity. The prevalence of achromatopsia has been estimated as 1:30,000 (for review, see [6, 7]), with approximately 20–30% of all cases caused by mutations in CNGA3 and 40–50% caused by mutations in CNGB3 [8]. In some instances, mutations in CNGA3 or CNGB3 result in cone dystrophy, a disease similar to achromatopsia but characterized by the progressive loss of cone function and sometimes progressive loss of rod function [9]. The steps that lead to rod or cone degeneration and, in particular, the role of CNG channel subunits in this process are not well understood. The recent success, however, in the identification of CNG channel mutants involved in visual diseases and the study of their defects by heterologous expression have provided insight into the molecular basis of photoreceptor dysfunction. The results of molecular genetic analyses of patients with

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Fig. A1. Mutations in cyclic nucleotide-gated (CNG) channel subunits associated with photo receptor dysfunction. See text for details. Underlined amino acid substitutions are found in patients with residual cone function. cNMP nucleoside 3′,5′-cyclic monophosphate, del deletion, Fs frame shift mutation, GARP glutamic acid-rich protein.

RP, achromatopsia, and cone dystrophy, as well as the in vitro experiments with the respective mutant CNG channel subunits are discussed next. Mutations in CNGA1 and CNGB1 Associated with Retinitis Pigmentosa Currently, six different mutant CNGA1 alleles have been identified in RP patients [1, 2] (Fig. A1). Three of these mutant alleles carry stop codons that terminate translation within the cytoplasmic N-terminus of CNGA1 (R28stop, E76stop, and K139stop). Another mutant CNGA1 allele is deleted in most of the protein-coding region and does not encode functional channels (not shown in figure). The remaining two mutant alleles encode either a single amino acid substitution (S316F) or an amino acid substitution and a truncation C-terminal to the cNMP-binding site (R654D-stop). Dryja and collaborators expressed CNGA1S316F and CNGA1R654D-stop in human embryonic kidney cells and found that these mutants form few functional homomeric channels [1]; cyclic guanosine monophosphate (cGMP)-dependent currents carried by single or few CNG channels were detected only in 3 of 85 (CNGA1S316F) or 1 of 83 membrane patches (CNGA1R654D-stop). Similar results were obtained with the bovine CNGA1R654D-stop homolog (CNGA1R656D-stop) expressed in Xenopus oocytes by Mallouk and collaborators [10]. Coexpression of CNGA1R656D-stop with CNGB1 did not rescue impaired channel

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trafficking. Surprisingly, neither truncation of the C-terminus (R656stop) nor the substitution R656D alone impaired surface expression. Apparently, the C-terminal five amino acids of CNGA1R656D-stop (KLKQD) generate a novel signal that causes retention in the endoplasmic reticulum (ER). Indeed, substitution of the C-terminal five amino acids of wild-type CNGA1 for KLKQD produces a mutant with impaired surface expression. Furthermore, Mallouk and coworkers provided evidence that the C-terminal sequence of CNGA1R656D-stop serves as a specific retention signal. Replacement of two key residues, leucine (L) or aspartate (D), in the C-terminal sequence KLKQD of CNGA1R656D-stop restored normal surface expression. This strong influence of single residues at the very C-terminus argues against the possibility that the mutations in CNGA1R656D-stop just cause improper protein folding. In contrast to the results reported by Dryja and Mallouk and their collaborators, Trudeau and Zagotta observed normal surface expression and channel formation of the human CNGA1 mutant CNGA1R654D-stop in Xenopus oocytes [11]. Impaired trafficking to the membrane occurs only when CNGA1R654D-stop is coexpressed with CNGB1. Using a biochemical pull-down assay, Trudeau and Zagotta demonstrated stable binding of a peptide representing the C-terminus of CNGA1 (amino acids 609–693) to a peptide that represents the region between the glutamic acid-rich protein (GARP) domain and the first transmembrane segment of CNGB1 (amino acids 677–764). Removing the C-terminal portion from the CNGA1 peptide, which corresponds to amino acids 657–693 in wild-type CNGA1, abolished this binding. Trudeau and Zagotta concluded from their results that the C-terminal amino acids 657–693 of CNGA1—which are deleted in R654D-stop—mask an ER retention signal within the amino acids 677–764 of CNGB1. In coexpression experiments with CNGB1 deletion mutants, this retention signal was confined to a segment of ten amino acids that precedes the first transmembrane segment of CNGB1 (YQFPQSIDPL). At present, it is not clear how the contradictory observations reported by Dryja and Mallouk and their collaborators on the one hand and by Trudeau and Zagotta on the other hand can be reconciled. Only a single mutation in the CNGB1 gene has yet been associated with RP [3]. The mutation results in the substitution of a highly conserved glycine for valine (G993V). This glycine is thought to reside in a turn between two β strands (β2 and β3) that contribute to a β-roll inside which the ligand binds [12]. The substitution of valine for glycine at this site might impair the relative orientation of the two β strands and result in a nonfunctional cNMP-binding site. Mutations in CNGA3 and CNGB3 Associated with Cone Dysfunction Genetic screens identified 57 mutant CNGA3 alleles that are associated with achromatopsia or cone dystrophy [9, 13–16]. The majority of these alleles (45) cause single amino acid substitutions, indicating that there is little tolerance for sequence variations in CNGA3 with respect to cone function. A genomic region that includes the CNGB3 gene was identified as a locus for achromatopsia in people from Pingelap, an atoll of Micronesia [17, 18]. About 5% of the Pingelap population is affected by achromatopsia, also known as Pingelapese blindness (OMIM 262300). This story gained notoriety as the subject of Oliver Sacks’ book, The Island of the Colorblind [19]. The prevalence of the disease traces back to 1775, when a typhoon decimated the population of Pingelap,

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Fig. A2. Achromatopsia affects color vision and visual acuity. A sunset as it might be seen by a person with normal color vision (A), by two sisters who confuse desaturated colors due to an incomplete form of achromatopsia (B), or by a person with complete achromatopsia (C).

leaving only a handful of survivors who repopulated the island. Genetic analysis identified a missense mutation in the CNGB3 gene (S435F) as the genetic basis of Pingelapese achromatopsia [20, 21]. Other than the Pingelap mutation, 15 additional mutant CNGB3 alleles have been associated with cone dysfunction [13, 15, 20–22]. Eight of these mutant alleles encode CNGB3 subunits lacking the cNMP-binding site due to stop-codon or frame-shift mutations. The remaining mutated CNGB3 alleles encode missense mutations in the p region or the cNMP-binding site, and one allele harbors a splice-site mutation in intron 13 (not shown in figure). Complete achromats are likely to carry mutant CNGA3 or CNGB3 alleles encoding nonfunctional channel proteins. Indeed, several mutant CNGA3 subunits associated with complete achromatopsia fold improperly and fail to reach the plasma membrane in heterologous expression systems [23, 24]. The fact that CNGA3 forms homomeric channels in heterologous expression systems raises the possibility that patients carrying CNGB3 null alleles do have cone CNG channels composed of CNGA3 only. In rod photoreceptors, however, the B subunit is essential for proper targeting of CNG channels to the outer segment [25]. Similarly, the lack of CNGB3 might cause low levels of CNGA3 in cone outer segments and thereby prevent cone function. In this respect, it would be informative to examine cone photoreceptors in dogs with naturally occurring mutations in CNGB3 [26]. In contrast to complete achromats, incomplete achromats are expected to carry at least one CNGA3 and one CNGB3 allele that allow some degree of cone function. Similarly, patients with cone dystrophy are likely to carry mutants that permit cone function until the cells degenerate. Alleles found exclusively in incomplete achromats or patients with cone dystrophy are underlined in Fig. A1. The in vitro characterization of CNGA3 or CNGB3 mutants encoded by these alleles might help scientists understand why the maintenance and degree of cone function varies among patients with impaired cone function. We analyzed the molecular basis of an incomplete form of achromatopsia, with considerable cone function, present in two sisters who both express the mutants CNGA3T224R and CNGA3T369S [27]. The sisters are able to discriminate saturated but not desaturated colors. Figure A2 gives an example of how the sisters would view colorful scenery. Detailed clinical characterization of the sisters, including psychophysical and electroretinographical analyses, showed that their cone system is characterized by

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lower light sensitivity and perturbed signal transfer from cones to postsynaptic neurons. Patch-clamp analysis of heterologously expressed mutant subunits revealed that only CNGA3T369S contributes to functional channels, although with altered ion permeation, gating, ligand sensitivity, and a weaker blockage by extracellular Ca2+ ions. Coexpression of wild-type CNGB3 with CNGA3T369S restored most of the native CNG channel properties except for the weaker Ca2+ blockage. The properties of CNGA3T369S/CNGB3 suggest that the mild form of achromatopsia in the sisters results from relatively subtle changes in ion flux through the cone CNG channel. Another channel defect appears to be common in patients with cone dystrophy. The CNGA3 mutations CNGA3N471S, CNGA3R563H, or the CNGB3 mutation CNGB3R403Q produce channels with slightly increased (less than twofold) cGMP sensitivity [28, 29]. This suggests that cone photoreceptors can cope with some increased CNG channel activity. This compensatory ability appears to be exhausted in the cones of Pingelap islanders. The Pingelap mutation CNGB3S435F produces channels with a roughly twofold increase in GMP sensitivity, invariantly leading to complete achromatopsia [22, 30]. Similarly, the mutations CNGB3D633G and CNGB3F525N, both associated with complete achromatopsia, confer an increase in cGMP sensitivity to CNG channels of two- and threefold, respectively [22, 29]. In conclusion, the degree of deviation from normal CNG channel function correlates with the degree of cone malfunction. Hence, we have to assume that photoreceptor function and survival critically depend on the correct tuning of CNG channel activation and permeability. With regard to future cures of photoreceptor dysfunction, the discovery of efficient CNG channel blockers (e.g., [31]) is particularly exciting. Substances that specifically block CNG channel currents might be used for the treatment of retinal diseases caused by abnormally high CNG channel activity. REFERENCES 1. Dryja, T. P., Finn, J. T., Peng, Y. W., McGee, T. L., Berson, E. L., Yau, K. W. (1995). Mutations in the gene encoding the alpha subunit of the rod cGMP-gated channel in autosomal recessive retinitis pigmentosa. Proc Natl Acad Sci U S A 92, 10177–10181. 2. Paloma, E., Martinez-Mir, A., Garcia-Sandoval, B., Ayuso, C., Vilageliu, L., GonzalezDuarte, R., Balcells, S. (2002). Novel homozygous mutation in the alpha subunit of the rod cGMP gated channel (CNGA1) in two Spanish sibs affected with autosomal recessive retinitis pigmentosa. J Med Genet 39, E66. 3. Bareil, C., Hamel, C. P., Delague, V., Arnaud, B., Demaille, J., Claustres, M. (2001). Segregation of a mutation in CNGB1 encoding the beta-subunit of the rod cGMP-gated channel in a family with autosomal recessive retinitis pigmentosa. Hum Genet 108, 328–334. 4. Pierce, E. A. (2001). Pathways to photoreceptor cell death in inherited retinal degenerations. Bioessays 23, 605–618. 5. Rattner, A., Sun, H., Nathans, J. (1999). Molecular genetics of human retinal disease. Annu Rev Genet 33, 89–131. 6. Sharpe, L. T., Nordby, K. (1990). Total Colorblindness: An Introduction. In: Night vision: basic clinical and applied aspects (Hess, R. F., Sharpe, L. T., Nordby, K., eds.), pp. 253. Cambridge University Press, Cambridge, UK. 7. Sharpe, L. T., Stockman, A., Jägle, H., Nathans, J. (1999). Opsin genes, cone photopigments, color vision and colorblindness. In: Color vision: from genes to perception (Gegenfurtner, K. R., Sharpe, L. T., eds.), p. 3. Cambridge University Press, Cambridge, UK.

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8. Kohl, S., Varsanyi, B., Antunes, G. A., Baumann, B., Hoyng, C. B., Jagle, H., Rosenberg, T., Kellner, U., Lorenz, B., Salati, R., Jurklies, B., Farkas, A., Andreasson, S., Weleber, R. G., Jacobson, S. G., Rudolph, G., Castellan, C., Dollfus, H., Legius, E., Anastasi, M., Bitoun, P., Lev, D., Sieving, P. A., Munier, F. L., Zrenner, E., Sharpe, L. T., Cremers, F. P., Wissinger, B. (2005). CNGB3 mutations account for 50% of all cases with autosomal recessive achromatopsia. Eur J Hum Genet 13, 302–308. 9. Wissinger, B., Gamer, D., Jagle, H., Giorda, R., Marx, T., Mayer, S., Tippmann, S., Broghammer, M., Jurklies, B., Rosenberg, T., Jacobson, S. G., Sener, E. C., Tatlipinar, S., Hoyng, C. B., Castellan, C., Bitoun, P., Andreasson, S., Rudolph, G., Kellner, U., Lorenz, B., Wolff, G., Verellen-Dumoulin, C., Schwartz, M., Cremers, F. P., Apfelstedt-Sylla, E., Zrenner, E., Salati, R., Sharpe, L. T., Kohl, S. (2001). CNGA3 mutations in hereditary cone photoreceptor disorders. Am J Hum Genet 69, 722–737. 10. Mallouk, N., Ildefonse, M., Pages, F., Ragno, M., Bennett, N. (2002). Basis for intracellular retention of a human mutant of the retinal rod channel alpha subunit. J Membr Biol 185, 129–136. 11. Trudeau, M. C., Zagotta, W. N. (2002). An intersubunit interaction regulates trafficking of rod cyclic nucleotide-gated channels and is disrupted in an inherited form of blindness. Neuron 34, 197–207. 12. Zagotta, W. N., Olivier, N. B., Black, K. D., Young, E. C., Olson, R., Gouaux, E. (2003). Structural basis for modulation and agonist specificity of HCN pacemaker channels. Nature 425, 200–205. 13. Johnson, S., Michaelides, M., Aligianis, I. A., Ainsworth, J. R., Mollon, J. D., Maher, E. R., Moore, A. T., Hunt, D. M. (2004). Achromatopsia caused by novel mutations in both CNGA3 and CNGB3. J Med Genet 41, e20. 14. Rojas, C. V., Maria, L. S., Santos, J. L., Cortes, F., Alliende, M. A. (2002). A frameshift insertion in the cone cyclic nucleotide gated cation channel causes complete achromatopsia in a consanguineous family from a rural isolate. Eur J Hum Genet 10, 638–642. 15. Nishiguchi, K. M., Sandberg, M. A., Gorji, N., Berson, E. L., Dryja, T. P. (2005). Cone cGMP-gated channel mutations and clinical findings in patients with achromatopsia, macular degeneration, and other hereditary cone diseases. Hum Mutat 25, 248–258. 16. Goto-Omoto, S., Hayashi, T., Gekka, T., Kubo, A., Takeuchi, T., Kitahara, K. (2006). Compound heterozygous CNGA3 mutations (R436W, L633P) in a Japanese patient with congenital achromatopsia. Vis Neurosci 23, 395–402. 17. Winick, J. D., Blundell, M. L., Galke, B. L., Salam, A. A., Leal, S. M., Karayiorgou, M. (1999). Homozygosity mapping of the Achromatopsia locus in the Pingelapese. Am J Hum Genet 64, 1679–1685. 18. Milunsky, A., Huang, X. L., Milunsky, J., DeStefano, A., Baldwin, C. T. (1999). A locus for autosomal recessive achromatopsia on human chromosome 8q. Clin Genet 56, 82–85. 19. Sacks, O. W. (1997). The island of the colorblind and Cycad Island. Knopf, New York. 20. Kohl, S., Baumann, B., Broghammer, M., Jagle, H., Sieving, P., Kellner, U., Spegal, R., Anastasi, M., Zrenner, E., Sharpe, L. T., Wissinger, B. (2000). Mutations in the CNGB3 gene encoding the beta-subunit of the cone photoreceptor cGMP-gated channel are responsible for achromatopsia (ACHM3) linked to chromosome 8q21. Hum Mol Genet 9, 2107–2116. 21. Sundin, O. H., Yang, J. M., Li, Y., Zhu, D., Hurd, J. N., Mitchell, T. N., Silva, E. D., Maumenee, I. H. (2000). Genetic basis of total colourblindness among the Pingelapese islanders. Nat Genet 25, 289–293. 22. Okada, A., Ueyama, H., Toyoda, F., Oda, S., Ding, W. G., Tanabe, S., Yamade, S., Matsuura, H., Ohkubo, I., Kani, K. (2004). Functional role of hCngb3 in regulation of human cone cng channel: effect of rod monochromacy-associated mutations in hCNGB3 on channel function. Invest Ophthalmol Vis Sci 45, 2324–2332.

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23. Faillace, M. P., Bernabeu, R. O., Korenbrot, J. I. (2004). Cellular processing of cone photo receptor cyclic GMP-gated ion channels: a role for the S4 structural motif. J Biol Chem 279, 22643–22653. 24. Patel, K. A., Bartoli, K. M., Fandino, R. A., Ngatchou, A. N., Woch, G., Carey, J., Tanaka, J. C. (2005). Transmembrane S1 mutations in CNGA3 from achromatopsia 2 patients cause loss of function and impaired cellular trafficking of the cone CNG channel. Invest Ophthalmol Vis Sci 46, 2282–2290. 25. Huttl, S., Michalakis, S., Seeliger, M., Luo, D. G., Acar, N., Geiger, H., Hudl, K., Mader, R., Haverkamp, S., Moser, M., Pfeifer, A., Gerstner, A., Yau, K. W., Biel, M. (2005). Impaired channel targeting and retinal degeneration in mice lacking the cyclic nucleotide-gated channel subunit CNGB1. J Neurosci 25, 130–138. 26. Sidjanin, D. J., Lowe, J. K., McElwee, J. L., Milne, B. S., Phippen, T. M., Sargan, D. R., Aguirre, G. D., Acland, G. M., Ostrander, E. A. (2002). Canine CNGB3 mutations establish cone degeneration as orthologous to the human achromatopsia locus ACHM3. Hum Mol Genet 11, 1823–1833. 27. Trankner, D., Jagle, H., Kohl, S., Apfelstedt-Sylla, E., Sharpe, L. T., Kaupp, U. B., Zrenner, E., Seifert, R., Wissinger, B. (2004). Molecular basis of an inherited form of incomplete achromatopsia. J Neurosci 24, 138–147. 28. Liu, C., Varnum, M. D. (2005). Functional consequences of progressive cone dystrophyassociated mutations in the human cone photoreceptor cyclic nucleotide-gated channel CNGA3 subunit. Am J Physiol Cell Physiol 289, C187–C198. 29. Bright, S. R., Brown, T. E., Varnum, M. D. (2005). Disease-associated mutations in CNGB3 produce gain of function alterations in cone cyclic nucleotide-gated channels. Mol Vis 11, 1141–1150. 30. Peng, C., Rich, E. D., Varnum, M. D. (2003). Achromatopsia-associated mutation in the human cone photoreceptor cyclic nucleotide-gated channel CNGB3 subunit alters the ligand sensitivity and pore properties of heteromeric channels. J Biol Chem 278, 34533–34540. 31. Brown, R. L., Haley, T. L., West, K. A., Crabb, J. W. (1999). Pseudechetoxin: a peptide blocker of cyclic nucleotide-gated ion channels. Proc Natl Acad Sci U S A 96, 754–759.

11 Rhodopsins in Drosophila Color Vision David Jukam, Preet Lidder, and Claude Desplan CONTENTS Introduction Anatomy and Molecular Aspects of Color-Sensitive Opsins in the DROSOPHILA Eye Development and Patterning of Rhodopsins for DROSOPHILA Color Vision Comparison Between Mammalian and DROSOPHILA Color Vision Rhodopsins Conclusion References

INTRODUCTION Color vision relies on an organism’s ability to detect color contrasts independently of intensity. This requires a sensory system that can detect and discriminate between different wavelengths of light. The opsin visual pigments collect light and initiate the conversion of photon energy into an electric signal sent by photoreceptor neurons to the brain, where this information is integrated into a perception. Opsins are photosensitive, seven-transmembrane G protein-coupled receptors of the same protein superfamily as many neurotransmitter, hormone, and sensory receptors that respond to molecular ligands [1, 2]. Different opsins exhibit different spectral sensitivities, and each is maximally sensitive to a particular wavelength of light, depending on its protein structure. Opsins are covalently linked to a retinal chromophore via a protonated Schiff base bond to a lysine residue in the seventh transmembrane domain of the protein [3, 4]. When light hits the opsin–chromophore complex (called rhodopsin), the 11-cis retinal chromophore isomerizes to all-trans retinal, leading to the activation of a G protein and downstream effectors. For example, in Drosophila melanogaster, a Gqα protein mediates the light-dependent activation of phospholipase C and opening of Transient Receptor Potential-Like (TRP/ TRPL) channels, resulting in the depolarization of the cell membrane [5–10]. Vertebrate and invertebrate rhodopsins act on different signal transduction cascades: Light depolarizes photoreceptors in insects and hyperpolarizes photoreceptors in vertebrates, but the

From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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end result of an activated rhodopsin is the same—a neural signal from the photoreceptor axon (invertebrates) or downstream neurons (retinal ganglion cells in vertebrates). The fruit fly Drosophila melanogaster is an excellent model for studying color vision because it has a sophisticated “hardware” to detect colors (including four different color-sensitive opsins) and can functionally and behaviorally respond to colors [11, 12, S. Yamaguchi and C. Desplan, unpublished results]. Although distantly related, flies and vertebrate opsins share the same biophysical properties, and Drosophila’s genetic malleability has supported fundamental discoveries in eye development, photoreceptor differentiation, visual transduction, and visual information processing. ANATOMY AND MOLECULAR ASPECTS OF COLOR-SENSITIVE OPSINS IN THE DROSOPHILA EYE Structure of the Drosophila Eye: Ommatidia, Photoreceptors, and Rhodopsins The Drosophila eye is organized into a latticed array of about 800 unit eyes, or ommatidia. Each ommatidium contains about 21 cells: 8 photoreceptors, 4 cone cells, a bristle cell, and about 8 accessory cells that help form the lens and pigment cells that shield photoreceptors (PRs) from light coming from other ommatidia [13]. The photoreceptors collect light in large, apical extensions of the membrane called rhabdomeres that contain densely packed rhodopsin molecules. The detection of light by rhodopsin in a photoreceptor initiates the signal phototransduction cascade that converts photon energy into an electrical impulse sent to higher-order neurons for visual information processing. According to their morphology, axonal projections, and opsin expression, the eight adult photoreceptors can be grouped into two main functional categories (Fig. 1): 1. The “outer” photoreceptors R1–R6 are specialized in dim-light vision and motion detection. Outer photoreceptors capture photons with high efficiency due to their broad-spectrum rhodopsin Rh1 as well as the large diameter and length of their

Fig. 1. Organization of the Drosophila eye. A Whole-eye view of a retina. Note the precise lattice of ommatidia. B Electron micrograph of a cross section through an ommatidium. The outer photoreceptors (R1–R6) are arranged in a chiral trapezoid, the center of which is occupied by the inner photoreceptors (R7 and R8), with R7 positioned on top of R8. C Longitudinal view of a single ommatidium. Light enters through the lens, at the top, and passes through the eight photoreceptor rhabdomeres.

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rhabdomeres, which extend from the basal to the apical side of the retina [14]. R1–R6 axons project to and terminate in the first optic ganglion, the lamina. They can be considered functionally analogous to vertebrate rods. These six outer photoreceptors are organized in a chiral trapezoid, which has a center that is occupied by the two “inner” photoreceptors, R7 and R8. 2. The “inner” photoreceptors R7 and R8 form the color detection system in the fly retina. The rhabdomeres of inner photoreceptors are in the same optical path, one beneath the other. R7 is located apically, closer to the lens, and expresses ultraviolet (UV)-sensitive Rh3 or Rh4. R8 is directly underneath R7 and expresses either blue-sensitive Rh5 or green-sensitive Rh6. R7 and R8 have thinner and shorter rhabdomeres than R1–R6, and their axons terminate in the second optic ganglion, the medulla [15]. Inner photoreceptors are similar to vertebrate cones in their role for color vision [16]. An additional layer of organization is imposed on the inner photoreceptors through the specific pairing of rhodopsins expressed in R7 and R8 (Fig. 2). R7 cells that express UV-sensitive Rh3 are always found in ommatidia with R8 cells that express the bluesensitive Rh5 [17, 18]. This Rh3/Rh5 rhodopsin expression in R7 and R8 photoreceptors defines the “pale” (p) ommatidial subtype. Conversely,R7 cells expressing a different UV-sensitive Rh4 are always found together with an R8 that expresses the green-sensitive Rh6, defining the “yellow” (y) ommatidial subtype [19, 20]. These two ommatidial subtypes are interspersed randomly within the fly retina [21], with 30% of the pale subtype and the remaining 70% of the yellow subtype. It is believed that the differences in rhodopsin between each subtype play a crucial role for the fly’s ability to discriminate between colors, with the p ommatidia discriminating among shorter wavelengths (UV to blue), and the y ommatidia discriminating longer wavelengths extending to the green. Thus, comparisons between wavelengths detected by R7 and R8 from the same ommatidium and comparisons among neighboring p and y R7/R8 subtypes are likely the first two steps in color vision processing. A third class of photoreceptors is located in the dorsal-most row of ommatidia (dorsal rim area, DRA) near the head cuticle. In these ommatidia, R7 and R8 both express Rh3 [22], which makes them inappropriate for color vision. Instead, the DRA inner photoreceptors detect the vector of oscillation plane of polarized light for spatial navigation [23, 24]. A fourth class of ommatidia, which does not exist in Drosophila, is found in the male Musca domestica in a large region of the eye where R7 expresses a Rh1-like molecule and projects to the lamina part of the optic lobe, thus leading to ommatidia that have seven motion detection photoreceptors instead of six and no color vision in this part of the retina [20]. This was entitled the “love spot” because it is used by males to track females during in-flight mating behavior. Because the DRA and Musca love spot are not involved in color vision, they are not discussed further. Molecular Genetics and Evolution of Rh5 and Rh6 The cloning of the gene encoding Rh5 [17, 18] provided the first opportunity to examine R8 opsin expression and to investigate color sensitivity in Drosophila. Introduction of either Rh5 or Rh6 into mutant strains that lack Rh1 expression is capable of restoring light response, indicating that these genes encode functional rhodopsins [25]. Flies expressing Rh5

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Fig. 2. Retinal mosaic in Drosophila. A Two main ommatidial subtypes can be distinguished: “pale” (p) and “yellow” (y). The model for p/y specification includes two steps: First, R7 cells choose between p and y fates by the stochastic expression of a transcription factor. Consequently, R7 cells impose their subset choice onto the underlying R8 cells (instruction). B Dorsal rim area ommatidia are always found in one or two rows at the dorsal periphery of the adult retina, whereas p (Rh3, Rh5) and y (Rh4, Rh6) ommatidia are distributed randomly through the retina in a 30:70 ratio. eq equator.

show peak sensitivity to wavelengths of 437 nm, corresponding to blue light, whereas Rh6 flies have a prominent peak at 508 nm, or green light (Table 1 [25]). The Rh5 protein shares several structural features with other insect opsins, such as the seven-transmembrane domains, an N-glycosylation site required for protein maturation [26], two cysteine residues that form a disulfide bond, and serine and threonine phosphorylation sites in the cytoplasmic C-terminal domain. In addition, Rh5 contains a tyrosine that is thought to serve as a counterion for the protonated Schiff base in the retinal chromophore and may affect light sensitivity.

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The gene encoding Rh6 was cloned using a Calliphora complementary DNA with significant homology to previously characterized fly rhodopsins [19]. Rh6, like Rh5, shares features consistent with the hallmarks of known rhodopsin molecules. The chromophore linkage sites, an N-glycosylation site, and cysteine residues to form a disulfide bridge are all present in Rh6, which also contains conserved proline residues in the transmembrane domain and the cytoplasmic domains responsible for G protein binding. Rh5 belongs to the blue family of invertebrate opsins and shares the highest amino acid similarity (59%) with locust S. gregaria lo2 opsin [18], also a blue opsin. Rh6 is a green opsin that is distant from the UV opsins. Other flies, such as Musca and the blowfly (Calliphora), which have a similar retinal organization as Drosophila, also have Rh5 and Rh6 orthologs [27]. Calliphora also has two R8 photoreceptor subtypes, pR8 and yR8, that are reported to have slightly different morphologies, a situation undetected in Drosophila melanogaster [28]. Drosophila Rh5 and Rh6 share higher amino acid identity with their Calliphora, honeybee, and mosquito orthologs than with other Drosophila rhodopsins, suggesting that rh5 and rh6 genes were both present in a common ancestor rather than having evolved independently in each fly lineage [27]. However, Rh5 is slightly more distant from the UV class—it shares 32% similarity with Rh3 and 33% with Rh4—than is Rh6, with 41% homology with Rh3 and 44% with Rh4 [17]. Together, this and other sequence evidence such as intron-exon structure, imply that Rh3/Rh4, Rh5, and Rh6 are ancestral rhodopsins, while a recent event in higher flies (after the mosquito lineage evolved) led to the duplication into the current Rh3 and Rh4. On the other hand, Rh5 is more divergent than Rh6 from Drosophila Rh1 (31%) and Rh2 (31%), which are themselves divergent members of the green class of invertebrate opsins (Fig. 3). Rh1 and Rh2 appear unique to the higher dipteran lineage and are replaced by several green-type opsins in bees and butterflies. They may have evolved from an ancestral Rh6-like green opsin and have evolved to acquire a very broad spectrum of absorption through the acquisition of a second retinal chromosphore that absorbs blue light and transfers it to the retinal inside the rhodopsin pocket [23]. Their evolution is likely linked to the appearance of an “open rhabdome” in these flies: The separation between the rhabdomeres of the different photoreceptors allowed for the utilization of a very broad-spectrum opsin such as Rh1.

DEVELOPMENT AND PATTERNING OF RHODOPSINS FOR DROSOPHILA COLOR VISION Mutually Exclusive Rhodopsin Expression Only after the Drosophila color-sensitive opsin genes had been cloned and protein localization determined could their genes and promoters be manipulated using the powerful genetic tools available in Drosophila. This allowed for the discovery of new genetic pathways and mechanisms that coordinate the complex expression pattern of rhodopsins in the fly retina. The “one-receptor molecule per neuron” rule in sensory systems ensures that the system can discriminate among sensory inputs by allowing each neuron to act as a

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Fig. 3. Phylogenetic relationship among color-sensitive rhodopsins. The phylogenetic tree displays the broad evolutionary relationship among opsin sequences. Vertebrate and invertebrate color-sensitive opsins likely evolved from a single common ancestor and later events created the subfamilies of green-, red-, blue-, and ultraviolet (UV)-sensitive opsins. GF = goldfish; Rh rhodopsin; ZF = zebrafish.

specialized unit of perception [29–33]. In the fly retina, photoreceptors also express “one rhodopsin per photoreceptor” to compare responses to different light wavelengths and create color contrast. In the Drosophila eye, the decision to express one of four rhodopsin molecules in inner photoreceptors, Rh3 or Rh4 in R7 and Rh5 or Rh6 in R8, establishes the basis for the retinal mosaic of photoreceptor subtypes and hence the basis for color vision [34]. It should be noted that coexpression of different opsins exists in bees and butterflies, although its significance is not clear and might correspond to the lack of the broad-spectrum Rh1 in these species. Even in mice, green and blue opsins are often found in the same cones, perhaps reflecting the fact that mice are nocturnal animals and practically color-blind. Although the distribution of p and y ommatidia is random, the 70:30 y:p ratio is constant from retina to retina across a large number of flies that diverged over 120 million years ago and rarely deviates more than 5% in either direction (35, 36, D. Vasiliauskas and C. Desplan, personal communication). This implies that photoreceptor subtype specification and color-sensitive rhodopsin expression are tightly regulated during development.

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The known mechanism for how photoreceptor subtypes are specified is, so far, one of the most detailed descriptions of how neurons express sensory receptors in a mutually exclusive manner. Next, we discuss recent results to illustrate the use of seemingly baroque and diverse mechanisms to control neuronal fate specification, rhodopsin expression, and retinal patterning, all resulting in one receptor per neuron. Transcription Factors Specify Outer from Inner Photoreceptors and Distinguish R7 from R8 The first step leading to mutually exclusive rhodopsin expression is the specification of a photoreceptor as belonging to the inner versus outer class. Eye development starts at the morphogenetic furrow, a wave of determination that moves across the developing eye imaginal disk during the third-instar larva life and leaves in its wake regularly spaced clusters of eight photoreceptors that will become each ommatidium. R8 is the first photoreceptor to be recruited, and it will recruit neighboring cells to become pairs of photoreceptors (R2/R5, R3/R4, and R1/R6) through sequential induction by the epidermal growth factor (EGF) receptor pathway. R7 is the last cell to be recruited, by a process that requires a dedicated signaling pathway, the Sevenless receptor tyrosine kinase and its ligand Boss [13]. It takes at least 4 days after R8 is first determined for R8 to complete its pale or yellow subtype specification, then differentiate and express Rh5 or Rh6. Which events in photoreceptor determination occur between the larval stage and the terminal differentiation in late pupal life, when all photoreceptors and subtypes are present? Cell–cell signaling and cell-specific transcription factors initially determine the identity of photoreceptors as outer or inner [37, 38]. The spalt gene complex (sal) directs R7 and R8 toward an inner photoreceptor state away from an outer photoreceptor “default” state. In sal mutants, all photoreceptors develop as “outer,” and all express Rh1. Once R7 and R8 are specified as generic inner photoreceptors, they continue toward more restricted fates. Further distinctions between R7 and R8 are achieved by the R7-specific expression of the transcription factor prospero, which is required to establish R7 fate and to repress R8 features (in particular Rh5 and Rh6 expression). A parallel role is ascribed to the zinc finger protein Senseless in R8 [39, 40]. The analysis of R7- and R8-specific rhodopsin promoters provides initial insight into mutually exclusive rhodopsin expression. Transgenic analysis shows that minimal promoters, often less than 300 bp in length, are sufficient to recapitulate Rh3, Rh4, Rh5, and Rh6 expression in vivo [39, 41, 42]. Although in Old World monkeys the exclusion mechanism between the M (green) and L (red) opsins is due to the regulation by a locus control region (LCR) distally located upstream of both genes, a similar mechanism can be ruled out in Drosophila because the four color rhodopsin genes in flies are not clustered and can be on distinct chromosomes [43–45]. Furthermore, even rhodopsin promoter transgenes are regulated appropriately and are expressed in the same exclusive subset as the endogenous gene with activity that they report, indicating trans-regulatory mechanisms. All four rhodopsin promoters contain a common rhodopsin conserved sequence 1 (RCSI)/palindromic 3 (P3) site near the TATA box; this site has been shown to provide general eye or photoreceptor-specific expression [46]. The site is recognized

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by the homeodomain of Pax-6, the “master control gene for eye development.” This function as a regulator of photoreceptor determination likely represents the ancestral role of Pax-6. Upstream from the RCSI/P3 site are rhodopsin-specific sequence elements common to the rhodopsins expressed in each photoreceptor (R7 vs. R8) or in each subtype (y or p). These appear to confer R7, R8, as well as subtype specific regulation. For instance, a conserved 11-bp element, seq56, found in both the rh5 and rh6 promoters is recognized by the R7-specific transcription factor Prospero. Binding by Prospero protein to seq56 in R7 prevents the expression of these R8 rhodopsins in R7 [39]. Indeed, when prospero or when the binding sites are mutated, Rh5 and Rh6 are expressed inappropriately in R7. Thus, Prospero restricts R8 rhodopsins to their proper cell type, R8. Another highly conserved site is found in the promoter of pR7 and pR8 rhodopsins, rh3 and rh5, respectively, which are always found in the same ommatidium. The homeodomain transcription factor Orthodenticle (Otd) binds to these sites, which are required for the p subtype: Rh3 and Rh5 are no longer expressed in otd eye mutants [42]. However, otd has only a permissive function as it is expressed in all photoreceptors, and its overexpression leads to no effect on rhodopsin transcription; other transcription factors must be required to control the pale R7/R8 subtype. Furthermore, the y pair, Rh4 and Rh6, is not expanded in otd mutants that lack the pR7 and pR8. Therefore, while the analysis of the promoters of Drosophila rhodopsins demonstrated that their mutually exclusive and subtype-specific rhodopsin regulation was transcriptional in nature (as opposed to posttranscriptional regulation at the messenger RNA or protein-trafficking level), genetics was used to unravel the mechanism of rhodopsin exclusion in Drosophila color vision. A Stochastic Decision Induces Rhodopsins in R7 Photoreceptor The specification to either the p or y ommatidial subtype is a two-step process that begins in R7 (Fig. 2). The instructive event that determines the 30:70 ratio of rhodopsins results from the stochastic expression of spineless (ss), which encodes yet another transcription factor, this time from the PAS-bHLH (per-arnt-sim, basic helix-loop-helix) family [46]. Only 70% of R7 cells express ss, and Rh4 is later induced in these same cells when they differentiate. The underlying R8 cells then turn on Rh6, which appears to be the default R8 opsin [47]. In those R7 that do not express ss, Rh3 is later induced as the R7 default state. However, these Rh3-expressing cells then instruct their R8 neighbor to express RH5 (see Fig. 4). It appears that the presence of ss causes Rh4 expression and prevents R7 from sending a signal to R8, whereas the absence of ss allows the R7-to-R8 signal to occur. Thus, ss is necessary and sufficient for the yR7 (Rh4, Rh6) subtype fate. The stochastic expression of spineless in a subset of R7 photoreceptors is the primary event that leads to the creation of the retinal mosaic of rhodopsin expression used for color vision. A Bistable Feedback Loop Specifies R8 Photoreceptor Subtype and Expression of Rh5 and Rh6 Strict pairing of rhodopsins in p and y R7/R8 subtypes could result from either a genetic program intrinsic to a particular ommatidium or from signaling events between

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Fig. 4. Mutually exclusive expression of color-sensing rhodopsins in R8. Schematic model for how the double-negative feedback loop directs rhodopsin expression in R8 photoreceptor. When R8 receives the signal from Rh3 expressing R7, the growth gene melted is expressed and suppresses warts, allowing for expressing of Rh5. If the signal is not received in R8, the tumor suppressor warts is expressed, which represses melted and promotes expression of Rh6. Thick black lines represent occurring genetic actions; thin gray lines represent genetic actions not occurring [48, 52]. Rh rhodopsin, ss spineless.

R7 and R8 that coordinate subtype specification. When R7 cells are genetically removed in sevenless mutant flies, all R8 express Rh6 [17, 18], suggesting that an R7 cell is required for Rh5 but not for Rh6 expression in R8. An R7-to-R8 signal model was thus proposed to explain the coordination between the rhodopsins in R7 and R8. How does R8 interpret the R7 signal and select its subtype as defined by expression of Rh5 or Rh6? While the identity of this signal remains elusive, genetic experiments have implicated the tumor suppressor warts/D-lats and a growth regulator melted in the postmitotic specification of R8 subtype fate [48] (Fig. 4). Warts is a serine/threonine kinase and the Drosophila homolog of the human large tumor suppressor (LATS) tumor suppressor genes that function to coordinately regulate proliferation and apoptosis in developing tissues [49, 50]. Warts misexpression in all photoreceptors induces the expression of Rh6 in all and only R8 cells. Conversely, warts loss of function leads to expression of Rh5 in all R8 [48]. This strongly suggests that warts is necessary and sufficient for the yR8 fate and for Rh6 expression. Whereas warts limits growth, melted is involved in fat metabolism as a positive regulator of the insulin/TOR (target of rapamycin) signaling pathway [51]. In R8, it opposes warts function to control Rh5 expression. Misexpression of melted and melted mutant

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flies have the opposite phenotypes of warts: All melted mutant R8 express Rh6, and overexpression of melted leads to expression of Rh5 in all R8 [48]. So, melted is both necessary and sufficient for pR8 fate and Rh5 expression. This suggests that warts and melted control each subtype through their mutual transcriptional regulation; indeed, they are expressed in mutually exclusive subsets of R8 cells that correspond to the y and p subtypes, respectively [48]. Wherever melted is expressed in R8, rh5 is induced; similarly, warts is coexpressed with Rh6. Therefore, warts and melted repress each other in a transcriptionally bistable negative-feedback loop that ensures that R8 cells only make one choice of expressing Rh5 or Rh6. This bistable loop interprets the R7 signal and instructs R8 to make a robust and unambiguous subtype choice [52] (Fig. 4). It should be noted that, although all other regulatory factors mentioned so far were transcription factors, Warts and Melted are cytoplasmic signaling proteins (Ser/Thr kinase and PH (pleckstrin homology domain) domain proteins, respectively). This reflects their involvement in interpreting the signal from R7 to decide between two alternate fates from an otherwise equipotent precursor cell. It remains that transcriptional effectors are required within the loop to control the expression of warts and melted. The involvement of such growth regulators in photoreceptor differentiation is an excellent example of how a signal transduction cassette can be reused during development for an entirely different purpose: Long after the photoreceptors have exited the cell cycle, the warts tumor suppressor pathway is no longer required and is now available to regulate this totally different process of cell specification. It should be noted that the other transcription factors described—Spalt, Prospero, Senseless, Orthodenticle, and Spineless—all have very crucial and totally different functions earlier in development. Some of them even act earlier in photoreceptor specification, before these genes are reused for specifying photoreceptor subtypes. COMPARISON BETWEEN MAMMALIAN AND DROSOPHILA COLOR VISION RHODOPSINS What can we learn about color vision from integrating data on mammalian and Drosophila rhodopsins? The vertebrate and invertebrate visual systems for color vision are different and have likely evolved independently, even though they are both based on very old photoreceptor molecules, the rhodopsins, which can even be found in bacteria. However, given the about 950 million years of evolution that separate Drosophila and mammals, the basic principles of function, organization, and development are remarkably similar [53]. Human Color-Sensitive Opsins Humans have three cone cell populations that detect short (S, blue-sensitive), medium (M, green-sensitive), and long (L, red-sensitive) wavelengths by expressing differentially sensitive opsins [43, 54]. Only one rhodopsin is usually expressed per cone type, although as in rabbit or mice, some coexpression is observed [55]. Dim-light vision and motion detection use the broadly tuned rod rhodopsin, which is evolutionarily a green opsin, likely because this represents the middle of the visible spectrum of light.

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This rhodopsin is specific to rods, which represent the majority of the human retina, and comprise almost the entire retina in nocturnal animals. Drosophila outer photoreceptors, like rods, also comprise the majority of photoreceptors as motion is critical for all of the fly visual functions. Outer photoreceptors also express a broad-spectrum rhodopsin (Rh1) that is related to green invertebrate opsins. Vertebrate and invertebrate opsins only share one very distant common ancestor of unknown spectral sensitivity, and it is likely that green opsins have evolved independently in the two branches. Although human cones are concentrated in the fovea (other animals do not exhibit this foveal specialization), the fly color receptors are distributed throughout the retina. However, in both cases, S, M, or L cone and fly color photoreceptors are distributed in a random manner [56]. This is in contrast to the fish retina, which contains four types of color photoreceptors that are precisely ordered, likely because they live in a blurry environment where repetitive patterns are unlikely to interfere with perception by regular arrays. Vertebrate retinal progenitors are multipotent and undergo successive phases of competence during which they successfully give rise to retinal ganglion cells, then cones and the neurons and glial cells that form the first layer of neural processing in the retina. Rods are produced throughout most of these phases. This depends on the expression of transcription factors that successively specify each type of retinal cell fates. The integration of these intrinsic transcription factor decisions ensures that each cell type is properly specified. Many human retinopathies that result in altered opsin expression are associated with mutations in transcription factors that regulate the specification of rod- and cone-specific genetic programs—Pax-6, Crx, Nrl, Nr2e3, and Trβ2—which are discussed below. Photoreceptor and Rhodopsin Specification in Flies and Mammals: Parallel Themes As in flies, the identification of factors controlling retinal development have come both from dissecting the promoter of the final differentiation products, the rhodopsin (e.g., nrl and crx), and from the genetic analysis of patients or mice with retinopathy (crx, nr2E3, tr2Beta). This resulted in the identification of two factors—Pax-6 and Crx—that have revealed similarities between vertebrate and Drosophila retina development that would not have been predicted from the independent evolution of the two visual systems. Pax-6, the master control gene for eye development in flies, is also a critical eye determination gene in vertebrates. Pax-6 mutations in mammals result in an early block in eye development and diseases such as aniridia. The common use of Pax-6 in fly and vertebrate development was explained by the ancestral role of Pax-6 in regulating photoreceptor determination, which led to its recruitment into controlling most steps in retinal (and lens) development. A second parallel between flies and mammals exists between Crx and Drosophila Otd, although the similarities are more difficult to explain. Crx is an Otd/Otx homeodomain transcription factor that plays a major role in controlling most opsins in vertebrates. Crx overexpression in mice produces more rod-like photoreceptors, while reducing Crx function disrupts photoreceptor morphogenesis [57]. Mutations in Crx occur in several inherited human retinopathies, including Leber’s congenital amaurosis and cone-rod dystrophy 2 [58]. Thus, Crx helps regulate photoreceptor differentiation and opsin expression. A functional analogy can be made with its ortholog in flies, Otd, which controls not only rh3 and rh5 but is also required

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for the proper repression of rh6 in outer photoreceptors and for rhabdomere morphogenesis [42]. Therefore, in both mammals and flies, an Otd/Otx family transcription factor appears to repress the expression of color-sensitive opsins in dim-light and motion detection photoreceptors. The recruitment of Otd for this function might relate to its role is early specification of anterior segments, which include the eye in vertebrates and the ocular segments in flies. Photoreceptor and Rhodopsin Specification in Flies and Mammals: Different Mechanisms A second transcription factor that controls the distinction between rods and cones is the bZIP (basic leucine zipper domain) protein NRL (neural retina leucine zipper). NRL positively regulates rod specification, in part by negatively regulating the transcription of cone genes in rods [59]. When NRL is removed in nrl−/− mice, the number of colorsensing cones (S cones) increases, while rod function is completely lost. This occurs because presumptive rods transform into “rod-cone” intermediates that exhibit hybrid rod-cone morphology but function as S cones because they express S-cone opsins [59]. Missense mutations in the NRL gene are associated with autosomal dominant retinitis pigmentosa, which further confirms a role for NRL in human photoreceptor development [60]. The nuclear hormone receptor Nr2E3 has also been implicated in the transcriptional regulation of photoreceptor subtype-specific genes. In a mouse model for retinal degeneration (rd7 mouse), more S cones are again present, while the number of rods is substantially decreased [60]. Nr2e3 may act in presumptive rod photoreceptors already expressing NRL to both activate a subset of rod specific genes and repress cone genes, including color-sensitive S-opsin [61]. A fourth gene that regulates mammalian rhodopsin expression is the thyroid hormone receptor TRβ2. It acts not by directly regulating the rod-cone decision but by controlling cone subtypes (Ng). TRβ2 mouse knockouts and cell culture showed that TRβ2 activates M-opsin and inhibits S-opsin, likely by reading a gradient of thyroid hormone during development [62, 63]. There are no known orthologs of NRL or TRβ2 in flies, and the only Drosophila ortholog of Nr2e3 has not been examined for eye function. Reciprocally, the two genes warts and melted play a critical role in the fly retina for specification of color photoreceptor subtypes, but it is unknown if their mammalian orthologs are involved in rhodopsin expression or if their mutations can cause human retinal pathologies. It is tempting to make a broad analogy between the role of NRL or NR2E3 with that of Drosophila prospero, which keeps particular color-sensitive rhodopsins repressed while simultaneously directing other aspects of photoreceptor morphogenesis [39]. Yet, the way these mammalian genes specify photoreceptors and regulate rhodopsin expression ultimately reflects the large mechanistic differences expected from two independently evolved eye systems. CONCLUSION Much has been learned about the molecular mechanisms that underlie Drosophila color vision, especially regarding molecular properties and developmental patterning

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of Rh5 and Rh6. In addition to increasing our understanding of the molecular basis of color vision, these insights allow one to manipulate the expression of all Drosophila rhodopsins and genetically encode neural activity reporters or inactivate individual neurons in the fly brain. When combined, these experimental technologies provide powerful tools for research into exciting, but unknown, areas such as color-dependent behaviors, how inputs are computed to form color perceptions, and the neural circuits responsible for color processing. REFERENCES 1. Sakmar, T. P., Menon, S. T., Marin, E. P., Awad, E. S. (2002). Rhodopsin: insights from recent structural studies. Annu Rev Biophys Biomol Struct 31, 443–484. 2. Terakita, A. (2005). The opsins. Genome Biol 6, 213. 3. Bownds, D. (1967). Site of attachment of retinal in rhodopsin. Nature 216, 1178–1181. 4. Wang, J. K., McDowell, J. H., Hargrave, P. A. (1980). Site of attachment of 11-cis-retinal in bovine rhodopsin. Biochemistry 19, 5111–5117. 5. Fain, G. L., Lisman, J. E. (1981). Membrane conductances of photoreceptors. Prog Biophys Mol Biol 37, 91–147. 6. Scott, K., Becker, A., Sun, Y., Hardy, R., Zuker, C. (1995). Gq alpha protein function in vivo: genetic dissection of its role in photoreceptor cell physiology. Neuron 15, 919–927. 7. Montell, C. (1999). Visual transduction in Drosophila. Annu Rev Cell Dev Biol 15, 231–268. 8. O’Tousa, J. E., Baehr, W., Martin, R. L., Hirsh, J., Pak, W. L., Applebury, M. L. (1985). The Drosophila ninaE gene encodes an opsin. Cell 40, 839–850. 9. Pak, W. L., Leung, H. T. (2003). Genetic approaches to visual transduction in Drosophila melanogaster. Receptors Channels 9, 149–167. 10. Yarfitz, S., Hurley, J. B. (1994). Transduction mechanisms of vertebrate and invertebrate photoreceptors. J Biol Chem 269, 14329–14332. 11. Tang, S., Guo, A. (2001). Choice behavior of Drosophila facing contradictory visual cues. Science 294, 1543–1547. 12. Quinn, W. G., Harris, W. A., Benzer, S. (1974). Conditioned behavior in Drosophila melanogaster. Proc Natl Acad Sci U S A 71, 708–712. 13. Wolff, T., Ready, D. F. (1991). The beginning of pattern formation in the Drosophila compound eye: the morphogenetic furrow and the second mitotic wave. Development 113, 841–850. 14. Zuker, C. S., Cowman, A. F., Rubin, G. M. (1985). Isolation and structure of a rhodopsin gene from D. melanogaster. Cell 40, 851–858. 15. Morante, J., Desplan, C. (2004). Building a projection map for photoreceptor neurons in the Drosophila optic lobes. Semin Cell Dev Biol 15, 137–143. 16. Kirschfeld, K., Franceschini, N. (1968). [Optical characteristics of ommatidia in the complex eye of Musca]. Kybernetik 5, 47–52. 17. Chou, W. H., Hall, K. J., Wilson, D. B., Wideman, C. L., Townson, S. M., Chadwell, L. V., Britt, S. G. (1996). Identification of a novel Drosophila opsin reveals specific patterning of the R7 and R8 photoreceptor cells. Neuron 17, 1101–1115. 18. Papatsenko, D., Sheng, G., Desplan, C. (1997). A new rhodopsin in R8 photoreceptors of Drosophila: evidence for coordinate expression with Rh3 in R7 cells. Development 124, 1665–1673. 19. Huber, A., Schulz, S., Bentrop, J., Groell, C., Wolfrum, U., Paulsen, R. (1997). Molecular cloning of Drosophila Rh6 rhodopsin: the visual pigment of a subset of R8 photoreceptor cells. FEBS Lett 406, 6–10.

264

Jukam et al.

20. Franceschini, N., Kirschfeld, K., Minke, B. (1981). Fluorescence of photoreceptor cells observed in vivo. Science 213, 1264–1267. 21. Franceschini, N. (1985). Early processing of colour and motion in a mosaic visual system. Neurosci Res Suppl 2, S17–S49. 22. Wernet, M. F., Labhart, T., Baumann, F., Mazzoni, E. O., Pichaud, F., Desplan, C. (2003). Homothorax switches function of Drosophila photoreceptors from color to polarized light sensors. Cell 115, 267–279. 23. Hardie, R. (1985). Functional organization of the fly retina (ed. Ottoson, D.) Prog Sens Physiol 5, 1–79 (Springer) 24. Labhart, T., Meyer, E. P. (2002). Neural mechanisms in insect navigation: polarization compass and odometer. Curr Opin Neurobiol 12, 707–714. 25. Salcedo, E., Huber, A., Henrich, S., Chadwell, L. V., Chou, W. H., Paulsen, R., Britt, S. G. (1999). Blue- and green-absorbing visual pigments of Drosophila: ectopic expression and physiological characterization of the R8 photoreceptor cell-specific Rh5 and Rh6 rhodopsins. J Neurosci 19, 10716–10726. 26. O’Tousa, J. E. (1992). Requirement of N-linked glycosylation site in Drosophila rhodopsin. Vis Neurosci 8, 385–390. 27. Schmitt, A., Vogt, A., Friedmann, K., Paulsen, R., Huber, A. (2005). Rhodopsin patterning in central photoreceptor cells of the blowfly Calliphora vicina: cloning and characterization of Calliphora rhodopsins Rh3, Rh5 and Rh6. J Exp Biol 208, 1247–1256. 28. Meffert, P., Smola, U. (1976). Electrophysiological measurements of spectral sensitivity of central visual cells in eye of blowfly. Nature 260, 342–344. 29. Mombaerts, P. (2004). Odorant receptor gene choice in olfactory sensory neurons: the one receptor-one neuron hypothesis revisited. Curr Opin Neurobiol 14, 31–36. 30. Vosshall, L. B., Amrein, H., Morozov, P. S., Rzhetsky, A., Axel, R. (1999). A spatial map of olfactory receptor expression in the Drosophila antenna. Cell 96, 725–736. 31. Clyne, P. J., Warr, C. G., Freeman, M. R., Lessing, D., Kim, J., Carlson, J. R. (1999). A novel family of divergent seven-transmembrane proteins: candidate odorant receptors in Drosophila. Neuron 22, 327–338. 32. Mazzoni, E. O., Desplan, C., Celik, A. (2004). “One receptor” rules in sensory neurons. Dev Neurosci 26, 388–395. 33. Serizawa, S., Miyamichi, K., Nakatani, H., Suzuki, M., Saito, M., Yoshihara, Y., Sakano, H. (2003). Negative feedback regulation ensures the one receptor-one olfactory neuron rule in mouse. Science 302, 2088–2094. 34. Wernet, M. F., Desplan, C. (2004). Building a retinal mosaic: cell-fate decision in the fly eye. Trends Cell Biol 14, 576–584. 35. Kirschfeld, K., Franceschini, N. (1977). Photostable pigments within the membrane of photoreceptors and their possible role. Biophys Struct Mech 3, 191–194. 36. Pichaud, F., Treisman, J., Desplan, C. (2001). Reinventing a common strategy for patterning the eye. Cell 105, 9–12. 37. Mollereau, B., Dominguez, M., Webel, R., Colley, N. J., Keung, B., de Celis, J. F., Desplan, C. (2001). Two-step process for photoreceptor formation in Drosophila. Nature 412, 911–913. 38. Cook, T. (2003). Cell diversity in the retina: more than meets the eye. Bioessays 25, 921–925. 39. Cook, T., Pichaud, F., Sonneville, R., Papatsenko, D., Desplan, C. (2003). Distinction between color photoreceptor cell fates is controlled by Prospero in Drosophila. Dev Cell 4, 853–864. 40. Frankfort, B. J., Nolo, R., Zhang, Z., Bellen, H., Mardon, G. (2001). senseless repression of rough is required for R8 photoreceptor differentiation in the developing Drosophila eye. Neuron 32, 403–414.

Drosophila Color Vision

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41. Fortini, M. E., Rubin, G. M. (1990). Analysis of cis-acting requirements of the Rh3 and Rh4 genes reveals a bipartite organization to rhodopsin promoters in Drosophila melanogaster. Genes Dev 4, 444–463. 42. Tahayato, A., Sonneville, R., Pichaud, F., Wernet, M. F., Papatsenko, D., Beaufils, P., Cook, T., Desplan, C. (2003). Otd/Crx, a dual regulator for the specification of ommatidia subtypes in the Drosophila retina. Dev Cell 5, 391–402. 43. Nathans, J., Merbs, S. L., Sung, C. H., Weitz, C. J., Wang, Y. (1992). Molecular genetics of human visual pigments. Annu Rev Genet 26, 403–424. 44. Smallwood, P. M., Wang, Y., Nathans, J. (2002). Role of a locus control region in the mutually exclusive expression of human red and green cone pigment genes. Proc Natl Acad Sci U S A 99, 1008–1011. 45. Nathans, J. (1999). The evolution and physiology of human color vision: insights from molecular genetic studies of visual pigments. Neuron 24, 299–312. 46. Sheng, G., Thouvenot, E., Schmucker, D., Wilson, D. S., Desplan, C. (1997). Direct regulation of rhodopsin 1 by Pax-6/eyeless in Drosophila: evidence for a conserved function in photoreceptors. Genes Dev 11, 1122–1131. 47. Wernet, M. F., Mazzoni, E. O., Celik, A., Duncan, D. M., Duncan, I., Desplan, C. (2006). Stochastic spineless expression creates the retinal mosaic for colour vision. Nature 440, 174–180. 48. Mikeladze-Dvali, T., Wernet, M. F., Pistillo, D., Mazzoni, E. O., Teleman, A. A., Chen, Y. W., Cohen, S., Desplan, C. (2005). The growth regulators warts/lats and melted interact in a bistable loop to specify opposite fates in Drosophila R8 photoreceptors. Cell 122, 775–787. 49. Justice, R. W., Zilian, O., Woods, D. F., Noll, M., Bryant, P. J. (1995). The Drosophila tumor suppressor gene warts encodes a homolog of human myotonic dystrophy kinase and is required for the control of cell shape and proliferation. Genes Dev 9, 534–546. 50. Xu, T., Wang, W., Zhang, S., Stewart, R. A., Yu, W. (1995). Identifying tumor suppressors in genetic mosaics: the Drosophila lats gene encodes a putative protein kinase. Development 121, 1053–1063. 51. Teleman, A. A., Chen, Y. W., Cohen, S. M. (2005). Drosophila Melted modulates FOXO and TOR activity. Dev Cell 9, 271–281. 52. Mikeladze-Dvali, T., Desplan, C., Pistillo, D. (2005). Flipping coins in the fly retina. Curr Top Dev Biol 69, 1–15. 53. Hedges, S. B. (2002). The origin and evolution of model organisms. Nat Rev Genet 3, 838–849. 54. Nathans, J., Thomas, D., Hogness, D. S. (1986). Molecular genetics of human color vision: the genes encoding blue, green, and red pigments. Science 232, 193–202. 55. Applebury, M. L. (1980). The primary processes of vision: a view from the experimental side. Photochem Photobiol 32, 425–431. 56. Roorda, A., Williams, D. R. (1999). The arrangement of the three cone classes in the living human eye. Nature 397, 520–522. 57. Furukawa, T., Morrow, E. M., Li, T., Davis, F. C., Cepko, C. L. (1999). Retinopathy and attenuated circadian entrainment in Crx-deficient mice. Nat Genet 23, 466–470. 58. Morrow, E. M., Furukawa, T., Raviola, E., Cepko, C. L. (2005). Synaptogenesis and outer segment formation are perturbed in the neural retina of Crx mutant mice. BMC Neurosci 6, 5. 59. Mears, A. J., Kondo, M., Swain, P. K., Takada, Y., Bush, R. A., Saunders, T. L., Sieving, P. A., Swaroop, A. (2001). Nrl is required for rod photoreceptor development. Nat Genet 29, 447–452. 60. Corbo, J. C., Cepko, C. L. (2005). A hybrid photoreceptor expressing both rod and cone genes in a mouse model of enhanced S-cone syndrome. PLoS Genet 1, e11.

266

Jukam et al.

61. Cheng, H., Aleman, T. S., Cideciyan, A. V., Khanna, R., Jacobson, S. G., Swaroop, A. (2006). In vivo function of the orphan nuclear receptor NR2E3 in establishing photoreceptor identity during mammalian retinal development. Hum Mol Genet 15, 2588–2602. 62. Yanagi, Y., Takezawa, S., Kato, S. (2002). Distinct functions of photoreceptor cell-specific nuclear receptor, thyroid hormone receptor beta2 and CRX in one photoreceptor development. Invest Ophthalmol Vis Sci 43, 3489–3494. 63. Harpavat, S., Cepko, C. L. (2003). Thyroid hormone and retinal development: an emerging field. Thyroid 13, 1013–1019.

12 INAD Signaling Complex of Drosophila Photoreceptors Armin Huber and Nina E. Meyer CONTENTS Introduction Comparison of Vertebrate and DROSOPHILA Phototransduction Cascades Identification of the INAD Signaling Complex Structure of the INAD Signaling Complex and Binding Specificity Anchoring of the INAD Signaling Complex to the Microvillar Membrane Function of the INAD Signaling Complex Information Transfer from Rhodopsin to the Signaling Complex BY the Visual G Protein Signaling Complexes in Vertebrate Photoreceptor Cells References

INTRODUCTION Phototransduction cascades convert the absorption of a photon into an electrical response of the photoreceptor cells by utilizing G protein-coupled signaling pathways (for reviews, see [1–5]). These pathways have been studied in great detail in photoreceptors of vertebrate and invertebrate model organisms such as bovine, mouse, toad, or Drosophila and squid. The work on these systems has provided much of the knowledge that we currently have about G protein-coupled signal transduction in general, and without doubt it will advance our understanding of these pathways in the future. To fulfill their function as highly efficient and fast light detectors, photoreceptors have optimized their signal transduction cascades toward high sensitivity and speed. Vertebrate as well as invertebrate photoreceptors are able to detect the absorption of a single photon [6, 7]. This requires an extremely high signal-to-noise ratio that can only be ensured when cross-talk from other signaling pathways is avoided and spontaneous activation of the phototransduction cascade is kept at a minimum. One strategy to avoid

From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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cross-talk is to physically separate the phototransduction cascade from other signaling pathways. In vertebrate photoreceptors, this is achieved by sequestering the phototransduction proteins into a specialized cellular compartment, the outer segment of rod or cone photoreceptors, that serves only for the detection of light and is connected to the rest of the cell by a small ciliary stalk. In Drosophila photoreceptors, proteins involved in light detection are also confined to a separate cell compartment, the rhabdomere formed by a densely packed stack of microvilli along one side of the cell. In addition to being confined to the rhabdomere, phototransduction proteins of Drosophila photoreceptors are organized into supramolecular complexes by the scaffolding protein INAD (inactivation no afterpotential D). The identification of the INAD signaling complex in these photoreceptor cells has led to a reevaluation of the principles by which Drosophila phototransduction is activated and regulated. With respect to signaling specificity, it has been suggested that the organization of the phototransduction cascade into signaling complexes avoids cross talk with other signaling cascades [8]. The concept of signaling complexes, in which signaling proteins are permanently linked together, differs fundamentally from a model in which signaling proteins diffuse freely in the photoreceptive membrane and interact with each other by random collisions. The model of freely diffusing signaling components has been established for vertebrate rod photoreceptor cells. In this sensory system, the high concentration of phototransduction proteins in the disk membrane may allow the cascade to operate at sufficient speed and specificity without forming signaling complexes, although some components of this signaling cascade appear to be linked together as well (see “Signalling complexes in vertebrate photoreceptor cells”). The compound eyes of fast-flying insects like Drosophila show a much higher temporal resolution than vertebrate eyes, and their photoreceptors have evolved the fastest G protein-coupled signaling cascades known to date. In Drosophila photoreceptors, the receptor potential can be generated within 20 ms after light stimulation [9]. This is about 10 times faster than in mammalian photoreceptors and about 100 times faster than in toad photoreceptors [3]. Arguably, the assembly of phototransduction proteins into signaling complexes may contribute to the high signaling speed of the cascade by eliminating delays due to the time required for protein diffusion. In this contribution, we first highlight common and discriminative features of the vertebrate and Drosophila phototransduction cascade. We then focus on the INAD signaling complex of Drosophila photoreceptors and finally discuss the assembly of signaling proteins in vertebrate photoreceptors. For a more comprehensive coverage of these topics, refer to excellent reviews on invertebrate photoreception in general [3, 4, 10–15] or on the INAD signaling complex in particular [8, 16–20]. COMPARISON OF VERTEBRATE AND DROSOPHILA PHOTOTRANSDUCTION CASCADES Photoreception in vertebrates and in Drosophila shows common features as well as significant differences. In both visual systems, light is absorbed by the seventransmembrane-receptor rhodopsin, which is present at high concentration in the disk membranes of rod photoreceptor cells or in the microvillar membranes of rhabdomeral photoreceptor cells. The high density of rhodopsin in the light-absorbing compartments

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ensures a high probability of catching a photon and hence a high sensitivity of the photoreceptors. Vertebrate and invertebrate rhodopsins display low but significant sequence identity, for example, 22% amino acid identity between the major Drosophila rhodopsin Rh1 and bovine rhodopsin [21, 22]. It is now generally accepted that all animal rhodopsins evolved from a common ancestor [23]. Conserved amino acids in vertebrate and invertebrate rhodopsins include a lysine in the seventh transmembrane segment that constitutes the chromophore-binding site, serine and threonine residues near the C-terminus that are targets for light-dependent rhodopsin phosphorylation, and consensus sites for N-linked glycosylation near the N-terminus of the protein. Although molecular details in rhodopsin activation and inactivation are markedly different between vertebrates and invertebrates [24, 25], the rhodopsin activation mechanism initiated by conversion of an 11-cis retinal chromophore into its all-trans form follows similar principles as does rhodopsin inactivation by binding of arrestin proteins to the active conformation of the receptor. To transmit the light signal from rhodopsin to photoreceptor ion channels, vertebrates and invertebrates make use of different kinds of G protein-coupled cascades (Fig. 1). Vertebrate rhodopsins activate the visual G protein transducin, which couples to a phosphodiesterase (Fig. 1A). Activation of the phosphodiesterase results in breakdown of cyclic guanosine monophosphate (cGMP), which is synthesized by a guanylate cyclase and, in the dark, keeps a fraction of cGMP-gated ion channels in an open state. Thus, light activation of most vertebrate photoreceptors leads to the closure of cGMP-gated ion channels, which results in a hyperpolarization of the cells. In Drosophila, as in most invertebrates, the activated rhodopsin molecule transmits the signal to a heterotrimeric Gq protein consisting of Gαq, Gβe, and Gγe (Fig. 1B). As a result of G protein activation, Gαq couples to the effector enzyme phospholipase Cβ (PLCβ). Activated PLCβ hydrolyzes the membrane phospholipid phosphatidylinositol 4,5-bisphosphate (PIP2) to form the second messengers diacylglycerol (DAG) and inositol 1,4,5-trisphosphate (IP3). The phototransduction cascade terminates in the opening of cation channels composed of ion channel subunits of the TRP protein family, TRP, TRPL, and possibly TRPγ, which results in the depolarization of the photoreceptor cell membrane. Although the gating mechanism of these TRP channels has not yet been entirely clarified, several lines of evidence suggest that the DAG branch, rather than the IP3 branch of the phosphoinositol signaling pathway activates the ion channels [26]. In both vertebrate and invertebrate photoreceptors, the absorption of a single photon results in the generation of a distinct electrophysiological response of the receptor, a so-called quantum bump. Quantum bumps correspond to the coordinated closure or opening of a defined number of ion channels. The macroscopic response to a stronger light stimulus (i.e., the receptor potential) is achieved by summation of the quantum bumps obtained from more or less simultaneous absorption of many photons. IDENTIFICATION OF THE INAD SIGNALING COMPLEX In the 1970s, Bill Pak and coworkers isolated a number of Drosophila phototransduction mutants that were identified by their altered electrophysiological characteristics in electroretinogram recordings [27, 28]. This approach eventually led to the identification of major players in Drosophila phototransduction such as rhodopsin Rh1 [21, 22] or the

Fig. 1. Comparison of vertebrate and Drosophila phototransduction cascades. A In vertebrate photoreceptors, the G protein transducin couples to a phosphodiesterase (PDE). The active PDE degrades cyclic guanosine monophosphate (cGMP), which is synthesized by a guanylate cyclase (GC) from guanosine triphosphate (GTP). In the dark, the cGMP-gated channels are kept open by cGMP. Light activation of these photoreceptors leads to the closure of the cGMPgated cation channels and thus to a hyperpolarization of the cell membrane. B In Drosophila photoreceptors, the Gq protein activates a phospholipase Cβ (PLCβ), which then hydrolyzes the membrane phospholipid phosphatidylinositol 4,5-bisphosphate (PIP2) and generates the two second messengers diacylglycerol (DAG) and inositol 1,4,5-trisphosphate (IP3). The phototransduction cascade terminates in the opening of at least two cation channels, TRP (transient receptor potential) and TRPL (TRP-like), which leads to a depolarization of the cell membrane. TRP and TRPL channels may be gated by DAG or by polyunsaturated fatty acids (PUFAs), which could be released from DAG by a DAG lipase. The scaffolding protein INAD (inactivation no afterpotential D) organizes phototransduction proteins into a supramolecular complex. The unconventional myosin NINAC (neither inactivation no afterpotential C) may couple this supramolecular complex to the actin cytoskeleton. An eye-specific protein kinase C (ePKC), which is also part of this complex, appears to be involved in response inactivation and light adaptation. Rh rhodopsin, SMC submicrovillar cisternae.

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central effector enzyme PLCβ [29]. Another mutant isolated in this genetic screen was termed inactivation no afterpotential D (inaD). The electroretinogram of this mutant showed inactivation by a strong light stimulus but lacked the afterpotential, a prolonged depolarization after termination of the light stimulus that is observed in wild-type photoreceptors when more than 20% of rhodopsin molecules are activated. The corresponding inaD gene was isolated by a subtractive hybridization screen for eye-enriched complementary DNAs, one of which rescued the inaD phenotype in transgenic Drosophila [30]. The inaD gene encodes a protein of 674 residues with homology to PDZ domain-containing proteins. PDZ domains have been named after the postsynaptic density protein PSD95, Drosophila disks large (Dlg), and the tight-junction protein ZO-1, in which this domain was first identified. Although initially only two PDZ domains were identified in INAD, Tsunoda et al. [31] realized that INAD is a pure scaffolding protein that is composed almost exclusively of five PDZ domains. As a first interaction partner of INAD, one of the two principle light-activated ion channels, TRP, was identified [32]. The binding of TRP to INAD is abolished in the original inaDP215 mutant by a point mutation (M442K) in the third PDZ domain. The existence of a signaling complex in fly photoreceptors was first proposed when coimmunoprecipitation studies with proteins of purified photoreceptor membranes from larger flies revealed that INAD, TRP, PLCβ, and an eye-specific protein kinase C (ePKC) are tethered together [33, 34]. The assembly of these phototransduction proteins was also shown for Drosophila photoreceptors, and a requirement of INAD for the rhabdomeral localization of TRP was demonstrated [35]. The concept of organizing major components of the fly visual transduction cascade into a signaling complex was further established and extended by Tsunoda et al. [31], who demonstrated that INAD functions as a scaffold to which the phototransduction proteins are bound. In a newly generated INAD-null mutant, in which the photoresponse was almost abolished, the signaling complexes were completely lost, and TRP, PLCβ, and ePKC were mislocalized. Besides the major INAD ligands TRP, PLCβ, and ePKC that are present in the complex at about equimolar ratios [33], additional photoreceptor proteins were shown to interact with INAD [35–37]. These proteins are rhodopsin, the second ion channel TRPL, the unconventional myosin NINAC (neither inactivation no afterpotential C), and calmodulin. They are detected in coimmunoprecipitation studies in substoichiometric concentrations, and their localization to the rhabdomere does not depend on the presence of INAD. While binding of the NINAC myosin III to INAD has been reported to be required for proper termination of the visual response [36], the significance of the proposed interaction of rhodopsin, TRPL, and calmodulin with INAD is not clear.

STRUCTURE OF THE INAD SIGNALING COMPLEX AND BINDING SPECIFICITY The three-dimensional structure of PDZ domains was first solved for the third PDZ domain of PSD95 and for the human homolog of the discs-large protein (hDlg) by x-ray diffraction of crystals containing the PDZ domains and short peptides corresponding to the C-terminal region of their targets [38, 39]. The approximately

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90 amino acids of these PDZ domains fold into a six-stranded β-sheet (B1–B6) and two α-helices (A1, A2). PDZ domains typically bind their targets by interaction with a short stretch of amino acids at the very C-terminal end of the proteins. The peptides bind in a groove between the B2 strand and the A2 helix and form an antiparallel β-strand that interacts with the B2 strand by hydrogen bonding. The region between A2 helix and B2 strand contains a conserved R/K-X-X-X-G-L-G-F motif that forms a carboxylatebinding loop that interacts with the free COO− group at the C-terminus of the ligand (for reviews, see [40, 41]). Screening of an oriented peptide library with nine different PDZ domains showed that consensus sequences for PDZ interaction fall into two main categories: (S/T)-XΦ-COOH and Φ-X-Φ-COOH (X is any amino acid and Φ is a hydrophobic amino acid) [42]. Accordingly, PDZ domains have been grouped into type I PDZ domains that bind (S/T)-X-Φ-COOH and type II PDZ domains that bind Φ-X-Φ-COOH. Additional binding motifs that do not fit into these categories have been identified, for example, the motif D/E-X-Φ-COOH in peptides binding to the PDZ domain of neuronal nitric oxide synthase [43] or H-W-C-COOH at the C-terminus of N-type Ca2+ channel that binds to the first PDZ domain of Mint-1 [44]. Due to this heterogeneity in binding motifs, it is not possible to predict from the amino acid sequence alone whether a given photoreceptor protein is a putative ligand for one of the PDZ domains of INAD. For the INAD signaling complex, structural details are only available for the binding of PLCβ to PDZ1 of INAD. The interaction of a heptapeptide corresponding to the C-terminal region of PLCβ with PDZ1 has been studied by crystallography [45]. The binding of this peptide to PDZ1 differs from typical PDZ–ligand binding schemes because it involves a covalent interaction. The crystal structure revealed a disulfide bond between the cysteine residue in the F-C-A-COOH motif of PLCβ and a cysteine in the B2 strand of the PDZ domain, which results in a high-affinity interaction between PLCβ and INAD. The functional significance of this covalent interaction is, however, not clear. The PLCβ homolog of the related fly species Calliphora has no cysteine near the C-terminus (A. Huber and M. Bähner, unpublished), and this PLCβ can be removed from INAD with high salt buffer under nonreducing conditions [33]. Studies using GST (glutathione S-transferase) pull-down assays, coimmunoprecipitation of recombinantly expressed PDZ domains with putative binding partners, yeast two-hybrid assays, and analysis of Drosophila mutants with mutations in specific PDZ domains or ligand-binding domains provided additional insights into the binding partners of each of the five PDZ domains of INAD. At least in vitro, it appears that binding of the various INAD ligands is not restricted to one particular PDZ domain for each target, for example, ePKC, which was originally shown to bind to PDZ4 was found to interact also with PDZ2 and PDZ3 [37, 46]. PLCβ was shown to bind to PDZ1 with its C-terminal binding motif F-C-A-COOH and also to PDZ5 with an internal sequence that overlaps with the G protein-binding domain [47]. In vivo, PLCβ binding to PDZ5 is relevant for PLCβ stability because a point mutation of a conserved glycine in PDZ5 or introduction of a stop codon between PDZ3 and PDZ4 that results in the expression of a truncated INAD significantly reduces the level of PLCβ [31, 48]. TRP that appears to be associated with PDZ3 in vivo [31, 32] was found to interact also with PDZ4 [37]. The additional possible binding partners of INAD, rhodopsin, TRPL, and the myosin III NINAC also interact

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Fig. 2. Model of proposed supercomplexes consisting of several INAD (inactivation no afterpotential D) signaling complex core units. The core units consist of four molecules INAD (shown as broad lines; PDZ domains are numbered 1–5), four TRP (transient receptor potential) subunits that form a functional cation channel, four molecules PLCβ (phospholipase Cβ), and four molecules ePKC (eye-specific protein kinase C). Possible protein–protein interactions that may contribute in the assembly of supercomplexes as discussed in the text are indicated by dotted lines.

with the PDZ domains of INAD in vitro, while the binding of calmodulin was reported to take place in the region between PDZ1 and PDZ2 [37]. Importantly, Xu et al. (1998) provided evidence that INAD molecules can bind to each other via interaction of PDZ domains 3 and 4. This raised the possibility that the INAD signaling core complexes are organized into larger units composed of several ion channels, phospholipases Cβ, and protein kinases C. These supercomplexes are referred to as signalplex or transducisome [16, 37]. A model for the possible organization of these supercomplexes is shown in Fig. 2. In addition to INAD–INAD interactions, the supercomplexes could be generated by the formation of tetrameric TRP ion channels from TRP momomeres of different INAD core complexes or by the simultaneous interaction of PLCβ with PDZ1 and PDZ5 of different INAD molecules. In addition, it has been shown that PLCβ forms homodimers that could tether together two INAD core complexes [45].

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ANCHORING OF THE INAD SIGNALING COMPLEX TO THE MICROVILLAR MEMBRANE Early studies on the INAD signaling complex have revealed that INAD is required for proper localization of its major ligands (TRP, ePKC, and PLCβ) to the rhabdomere as these signaling proteins are mislocalized in inaD mutants and become degraded [16, 31, 35]. But, how is INAD itself anchored to the microvillar membrane? The localization of the INAD signaling complex to the rhabdomere could be achieved by interaction with components of the microvillar membrane or with the microvillar cytoskeleton. Insight into this question came from findings showing that TRP is crucial for anchoring INAD to the rhabdomeral membrane [49, 50]. INAD becomes mislocalized in trp null mutants or in mutants lacking the PDZ-binding site of TRP but not in mutants defective in other INAD ligands. Together with INAD, ePKC and PLCβ are also mislocalized in a trp mutant, and they are still attached to INAD in this mutant, as revealed by coimmunoprecipitation. Since TRP is the only transmembrane protein among the major components of the INAD signaling complex, it seems reasonable to assume that TRP is required for membrane attachment of the complex. However, a substantial amount of the signaling complex (ca. 25%) of INAD [50] remains in the rhabdomeres of trp mutants, suggesting the existence of an additional TRP-independent mechanism for INAD anchorage. The signaling complexes remaining in the rhabdomere of trp mutants may account for the fact that the electrophysiological phenotype of a trp null mutant, in which the receptor potential is generated by TRPL channels, is by far not as severe as in the inaD null mutant, which shows almost no light response [31]. Expression of INAD proteins with mutations in either of its five PDZ domains in CHO (Chinese hamster ovary) cells indicated that PDZ1 is involved in attaching INAD to the cell membrane [50]. Besides its interaction with PLCβ, PDZ1 of INAD was shown to bind to the unconventional myosin III NINAC, which in Drosophila photoreceptors could anchor the signaling complex to the microvillar actin cytoskeleton [36]. More recently, an interaction of TRP (and of TRPL) with the actin-binding protein moesin has been reported, which could represent an additional mechanism for linking the signaling complex to the cytoskeleton [51]. This last interaction is light dependent: Moesin is bound to the ion channels in dark-raised flies and released after 1 h of light adaptation. The dynamic interaction with TRP and TRPL results in a subcellular redistribution of moesin and concomitant rearrangements of the actin cytoskeleton. The question whether the TRP-INAD interaction is necessary for targeting the signaling complex to the rhabdomere or whether TRP and INAD reach the rhabdomere independently of each other and then require the interaction for staying there has also been clarified. In late pupae, at a time when maturation of rhabdomeres is completed and signaling proteins are first synthesized, TRP was found in the rhabdomeres independently of INAD, and INAD likewise reached the rhabdomeres independently of TRP. These findings suggest that the TRP–INAD interaction is necessary for anchoring of the signaling complex but not for its targeting [49, 50]. On the other hand, in wild-type flies the assembly of at least three proteins of the signaling complex (INAD, ePKC, PLCβ) seems to occur before its components have reached the rhabdomere. Evidence for this conclusion comes from experiments, in which INAD is expressed in the photoreceptors under the control of a heat shock promoter. While recovery of the electrophysiological

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response, which indicates the arrival of functional signaling complexes at the rhabdomeral membrane, was first measured 7 h after the heat shock, interaction of the signaling proteins, as determined by coimmunoprecipitation, was detected already 3 h after the activation of INAD expression by heat shock [50]. The preassembly of the signaling complex could ensure that only complete complexes and no stray complex components are present in the rhabdomere. It may also be required for the correct targeting of ePKC and PLCβ to the rhabdomere and thereby reduce the problem of targeting several proteins to the same location to targeting a single unit. The stage at which TRP enters the signaling complex has not been clarified so far.

FUNCTION OF THE INAD SIGNALING COMPLEX As outlined, INAD determines the localization of the essential phototransduction proteins TRP, PLCβ, and ePKC in the microvillar membrane of the rhabdomeres. Concentration of these proteins in a subcellular signaling compartment specialized for phototransduction certainly enhances signaling specificity by avoiding cross talk with other signaling cascades operating in the photoreceptor cell. Whether the association into the macromolecular INAD signaling unit within the microvillar membrane adds further specificity to the phototransduction pathway is questionable because no signaling pathway other than the phototransduction cascade is known to operate in the rhabdomere of mature photoreceptor cells. Most of the rhodopsin molecules are strictly confined to the rhabdomere but are not part of the signaling complex and therefore could potentially cross talk to other pathways if these pathways were present at this subcellular site. The situation is even worse with respect to the visual G protein and the major visual arrestin (Arr2), which are distributed throughout the photoreceptor cell. Immunostaining with antibodies against the visual G protein subunits detected Gαq, Gβ, and Gγ not only in the rhabdomeres but also in the cell bodies and even in the axons of the photoreceptor cells projecting into the optic ganglia [52, 53]. Gαq was shown to translocate light dependently between the rhabdomere and the cell body, thereby reducing its amount in the rhabdomere in light-raised flies [54, 55]. This mechanism may result in long-term adaptation of the photoreceptor cell. Arr2 translocates in the opposite direction: A major fraction of Arr2 is only present in the rhabdomere when the flies are light adapted, whereas in the dark Arr2 is localized in the cell body [56]. These findings suggest that signaling specificity of the Drosophila phototransduction cascade is mainly determined by compartmentalization into the rhabdomere of the receptor rhodopsin, the effector PLCβ, one of the two principle light-activated ion channels, TRP, and the regulatory protein ePKC. Compartmentalization of the last three components is achieved by formation of the INAD signaling complex. The second light-activated ion channel, TRPL, also translocates to the cell body on prolonged illumination [57]. In addition to a function of the INAD signaling complex in signaling specificity, it has been suggested by several authors that the assembly of signaling proteins may enhance signaling speed by reducing or eliminating diffusional delays [16, 31, 33, 58]. The classical inaD mutant, inaDP215, which has a point mutation in the third PDZ domain and fails to bind TRP, displays a defect in response termination [30, 32]. It has been shown

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that this defect results primarily from prolongation of the latencies for quantum bump generation [31], although a change in the shape of the quantum bumps may contribute to the defect also [59]. In any case, these findings suggest that the interaction between TRP and INAD is required for fast kinetics of the photoresponse. However, this conclusion is contradicted by results obtained with mutants expressing a C-terminally truncated TRP channel, TRP∆1272, which lacks the INAD-binding site [49]. Electrophysiological analysis of flies expressing TRP∆1272 at young age, when the TRP channel is still localized to the rhabdomeres, revealed no obvious differences compared to wild-type flies. A more recent article that analyzed the rescue of retinal degeneration in the rdgA mutant by mutations in INAD showed that the constitutive activity of TRP channels observed in the rdgA mutant does not depend on the TRP–INAD interaction [48]. It is possible that the inaDP215 mutation affects the binding of ligands other than TRP, and that this defect rather than the disrupted interaction with TRP is responsible for prolonged response termination. More severe defects in the photoresponse have been described for mutations that disrupt the interaction between PLCβ and INAD. In inaD2 and norpAC1094S, which have a mutation in PDZ2 or in the INAD-binding site of PLCβ, respectively, light sensitivity is reduced, and response latency and kinetics are substantially prolonged [31, 58]. Finally, the linking of TRP and ePKC in the INAD signaling complex may be important for proper deactivation of the visual response. TRP is phosphorylated in vitro by ePKC [60, 61], and mutation of the ePKC phosphorylation site Ser982 to alanine leads to slow deactivation of the visual response similar to that of InaDP215 [62]. In conclusion, it appears that although TRP can be activated independently of INAD, the kinetics of the photoresponse do depend on the correct assembly of the INAD signaling complex or at least on the INAD-dependent recruitment of the signaling components to the rhabdomeral photoreceptive membrane. INFORMATION TRANSFER FROM RHODOPSIN TO THE SIGNALING COMPLEX BY THE VISUAL G PROTEIN In vertebrate as well as in fly photoreceptors rhodopsin is the most abundant protein of the photoreceptive membrane. For the photoreceptors of larger flies (Calliphora), it has been estimated that 3–5 Gαq molecules and the same number of tetrameric TRP channels are present per 100 molecules of rhodopsin [33]. Since the main components of the signaling complex are thought to be present at an equimolar ratio, each tetrameric TRP channel would be associated with four molecules of INAD, PLCβ, and ePKC. A single photoreceptor microvillus harbors about 1,200 rhodopsins, 50 Gαq molecules, and 50 signaling complex core units (each composed of 4 TRP, 4 PLCβ, 4 ePKC, and 4 INAD molecules). The major fraction of rhodopsin is not tethered to the INAD signaling complex. Assuming that all rhodopsins in a microvillus have the same probability to absorb a photon, the activated rhodopsin may be close or relatively far away from the signaling complex. The spatial distance between activated rhodopsin and the complex has to be bridged, a task that is likely to be the function of the visual G protein. In this scheme (Fig. 3), the Gαq subunit would transport the signal by diffusion from activated rhodopsin to the next-available signaling complex. Indeed, it has been shown that Gαq is not permanently associated with the signaling complex but

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Fig. 3. Organization of the phototransduction cascade in Drosophila. The membrane is densely packed with rhodopsin molecules (Rh; ca. 1,200 per microvillus) and several INAD (inactivation no afterpotential D) signaling complex core units are present. The distance between activated rhodopsin and the next INAD signaling complex is presumably bridged by diffusion of the alpha subunit of the visual G protein. Activation of one rhodopsin molecule results in the coordinated gating of, on average, 15 TRP channels, which may be assembled into a supercomplex. Only two of the four subunits of INAD, PLCβ (phospholipase Cβ), and ePKC (eye-specific protein kinase C) per TRP tetramer are shown. NINAC (neither inactivation no afterpotential C).

interacts with PLCβ subunits that are attached to INAD only after G protein activation [63]. The varying distances between an activated rhodopsin and the signaling complex may underlie the observed variability of latencies of single-photon responses, which amount to between 20 and 100 ms [59]. In other words, the latencies may mainly reflect the time required for the shuttling of Gαq from rhodopsin to the signaling complex. In vertebrate photoreceptors, amplification of the signal is achieved at several levels. In this cascade, one activated rhodopsin activates several hundred G proteins. Each G protein activates one phosphodiesterase that hydrolyzes thousands of cGMP molecules, resulting in closure of numerous cGMP-gated ion channels [1, 2]. In sharp contrast, the Drosophila phototransduction cascade shows relatively little amplification at the level of the G protein. Studies of single-photon responses using severe Gαq hypermorphs showed that a reduction of Gαq subunit to 1% of wild-type level had little effect on the size of the quantum bumps [64, 65]. Instead, the frequency at which quantum bumps were generated was decreased dramatically in the Gαq mutant, leading to a more than 1,000-fold reduction in light sensitivity. Interestingly, the latency of the single-photon responses was not altered in the Gαq mutant. These findings suggest that a single G protein that has encountered an activated rhodopsin triggers the single-photon response like an on/of switch [64]. The data also suggest that there is no amplification at the level of the G protein. A more recent analysis of quantum bumps in the Gαq mutant questioned this last conclusion and suggested instead that one activated rhodopsin may activate about five G proteins [66]. In any case, the amplification at this stage is still relatively little when compared to the situation in vertebrate photoreceptors. To achieve the high sensitivity that allows the detection of a single photon by vertebrate as well as by invertebrate photoreceptors, noise resulting from spontaneous activation of

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signaling components has to be minimized. For example, it has been calculated that spontaneous (i.e., light-independent) activation of a given rhodopsin molecule occurs only once in 3,000 years [67]. In vertebrate photoreceptors, in which one rhodopsin activates many G proteins, spontaneous activation of a single G protein may not contribute significantly to background noise. In Drosophila, however, spontaneous activation of one G protein molecule can result in a quantum bump. Although the mechanism is not fully understood, it has been shown that a 2.5-fold excess of the Gβγ subunit over Gαq is required to suppress spontaneous G protein activation in Drosophila photoreceptor cells [68]. The extra Gβγ is not present in the microvillar membrane, where equal amounts of Gβγ and Gαq were found, but it was detected in the soluble fraction. Yet, a reduction of the total amount of Gβ to 50% results in a dramatic increase in the frequency of spontaneous quantum bump generation. The interaction of Gαq with its effector enzyme PLCβ not only activates PLCβ, but it also turns on the guanosine triphosphatase (GTPase) activity of the Gα-subunit. Thus, in Drosophila phototransduction PLCβ acts as a GAP (GTPase activating protein) accelerates that Gαq inactivation [69]. In Drosophila mutants with a reduced amount of PLCβ, the latency of quantum bump generation is highly increased because, on average, it takes much longer before Gαq encounters a PLCβ molecule and becomes inactivated [65, 69]. At the macroscopic level, PLCβ hypomorphs display prolonged activation of the photoresponse. With respect to signal amplification, these results suggest that there is no amplification at this step of the transduction cascade; that is, one activated G protein activates just one PLCβ. In analogy to vertebrate photoreceptors, it has been assumed that the absorption of one photon affects the gating of a large number of light-sensitive ion channels. However, more recent calculations of single-channel conductance of the TRP channels and average size of quantum bumps showed that the simultaneous opening of about 15 channels is sufficient for generating the single-photon response [59]. However, on subsequent activation of (different) rhodopsin molecules, the quantum bump size shows great variability, suggesting that the number of TRP channels that open in response to the absorption of one photon varies as well. An interesting experiment that provides some insight into the relationship between activation of rhodopsin and the number of gated TRP channels was performed by Scott and Zuker [65]. The authors used a calmodulin mutant that has a defect in rhodopsin inactivation. In this mutant, the same rhodopsin activated the cascade again and again, producing a train of quantum bumps. Surprisingly, the variability in the size of these quantum bumps was much smaller than in quantum bumps generated after activation of different rhodopsin molecules. Assuming that the same rhodopsin always activates the same signaling complex, this finding led to the conclusion that a quantum bump represents the output of a localized signaling complex that contains a defined number of ion channels. Variability in quantum bump size would then be due to differences in the composition of individual signaling complexes [65]. SIGNALING COMPLEXES IN VERTEBRATE PHOTORECEPTOR CELLS The cGMP-gated cation channels are the targets of the vertebrate phototransduction cascade, and these channels are closed on light stimulation, which results in hyperpolarization of the photoreceptor membrane. The cGMP-gated ion channels consist of an

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α- and a β-subunit [70]. The β-subunit has a large cytosolic N-terminal domain, which is referred to as the GARP (glutamine acid-rich protein) region. In addition, rod photoreceptors contain two splice variants of soluble GARPs, GARP1 and GARP2. Similar to INAD in Drosophila photoreceptors, the GARP proteins of rod photoreceptors seem to form a scaffold for the assembly of signaling components of the vertebrate photoreceptor [71, 72]. The GARP proteins contain four proline-rich repeats, which serve as binding sites for ligands. The exact composition of the GARP signaling complex is still unclear. Körschen et al. [71] suggested that the α- and the β-subunits of the cGMP-gated ion channel, as well as the guanylate cyclase, the phosphodiesterase, and an ATP-binding cassette transporter (ABCR) interact with GARP. The phosphodiesterase binds to the complex in its activated form and is inhibited by GARP. In addition, the cytoskeletal elements tubulin and actin have been shown to associate with one of the proline-rich repeats of GARP. As the cGMP-gated channel is localized in the plasma membrane of rod outer segments, whereas guanylate cyclase, phosphodiesterase, and ABCR are associated with the disk membranes, the GARP signaling complex has been proposed to bridge the gap between plasma membrane and disk membrane. This hypothesis is in agreement with the subcellular localization of the GARP proteins at the rim region and incisures of disks. In the above cited in vitro study [71], synthetic peptides containing each of the four proline-containing repeat regions were coupled to Sepharose beads and used as an affinity matrix to identify GARP-binding proteins. Poetsch et al. [72] also investigated possible interactions of GARP proteins with various soluble and membrane-bound proteins of rod outer segments. They used highly specific monoclonal antibodies in conjunction with immunoprecipitation, cross-linking, and electrophoretic techniques to identify membrane proteins of the rod outer segment that interact with GARP proteins. They found that the cGMPgated ion channel and soluble GARP proteins specifically associate with peripherin 2-containing oligomeric complexes at the rim region of disk membranes but not with ABCR or guanylate cyclase. In addition, they show that the cGMP-gated ion channel interacts with the Na/Ca-K exchanger present as a dimer in the plasma membrane of the rod outer segment, which is in agreement with earlier studies [73–76]. GARPmediated protein–protein interactions between the channel-exchanger complex in the plasma membrane of rod outer segments and peripherin 2 complexes in the rim region of the disk membrane may play a role in maintaining the spatial arrangement of the disk and plasma membrane in rod outer segments and anchoring the channelexchanger complex in the plasma membrane, preventing it from freely diffusing in the membrane [72]. While the Drosophila INAD signaling complex may play a role in enhancing signaling speed and specificity, the GARP complexes of vertebrate rod photoreceptors have been proposed to function as “adaptational” signaling complexes by inhibiting light-dependently bound phosphodiesterase and thereby preventing high cGMP turnover during daylight, when rod function is saturated [71, 72]. Alternatively, soluble GARP proteins may serve to cap peripherin 2 complexes at the rim region, preventing the C-terminus of peripherin 2 from initiating fusion of the disks with the plasma membrane [72, 77].

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ACKNOWLEDGMENTS We wish to thank J. Pfannstiel and I. Huber for critical comments on the manuscript. Our work is supported by grants from the Deutsche Forschungsgemeinschaft (Hu 839/2-4), the German-Israel-Foundation (GIF), and the Hertie-Foundation. REFERENCES 1. Lamb, T. D. (1996). Gain and kinetics of activation in the G-protein cascade of phototransduction. Proc. Natl. Acad. Sci. U. S. A. 93, 566–570. 2. Stryer, L. (1986). Cyclic GMP cascade of vision. Annu. Rev. Neurosci. 9, 87–119. 3. Hardie, R. C., Raghu, P. (2001). Visual transduction in Drosophila. Nature 413, 186–193. 4. Montell, C. (1999). Visual transduction in Drosophila. Annu. Rev. Cell Dev. Biol. 15, 231–268. 5. Koutalos, Y., Yau, K. W. (1996). Regulation of sensitivity in vertebrate rod photoreceptors by calcium. Trends Neurosci. 19, 73–81. 6. Wu, C. F., Pak, W. L. (1975). Quantal basis of photoreceptor spectral sensitivity of Drosophila melanogaster. J. Gen. Physiol. 66, 149–168. 7. Baylor, D. A., Lamb, T. D., Yau, K. W. (1979). Responses of retinal rods to single photons. J. Physiol. 288, 613–634. 8. Tsunoda, S., Sierralta, J., Zuker, C. S. (1998). Specificity in signaling pathways: assembly into multimolecular signaling complexes. Curr. Opin. Genet. Dev. 8, 419–422. 9. Ranganathan, R., Harris, G. L., Stevens, C. F., Zuker, C. S. (1991). A Drosophila mutant defective in extracellular calcium-dependent photoreceptor deactivation and rapid desensitization. Nature 354, 230–232. 10. Zuker, C. S. (1996). The biology of vision of Drosophila. Proc. Natl. Acad. Sci. U. S. A. 93, 571–576. 11. Minke, B., Parnas, M. (2006). Insights on TRP channels from in vivo studies in Drosophila. Annu. Rev. Physiol. 68, 649–684. 12. Minke, B., Hardie, R. C. (2000). Genetic dissection of Drosophila phototransduction. In: Handbook of biological physics, Volume 3: molecular mechanisms in visual transduction (Stavenga, D., DeGrip, W. J., Pugh, E. N., Jr., eds.), pp. 449–525. Elsevier, Amsterdam. 13. Pak, W. L., Leung, H. T. (2003). Genetic approaches to visual transduction in Drosophila melanogaster. Receptors Channels 9, 149–167. 14. Minke, B., Cook, B. (2002). TRP channel proteins and signal transduction. Physiol. Rev. 82, 429–472. 15. Paulsen, R., Bähner, M., Huber, A., Schillo, M., Schulz, S., Wottrich, R., Bentrop, J. (2001). The molecular design of a visual cascade: Molecular stages of phototransduction in Drosophila. In: Vision: the approach of biophysics and neurosciences (Musio, C., ed.), pp. 41–59. World Scientific, Singapore. 16. Tsunoda, S., Zuker, C. S. (1999). The organization of INAD-signaling complexes by a multivalent PDZ domain protein in Drosophila photoreceptor cells ensures sensitivity and speed of signaling. Cell Calcium 26, 165–171. 17. Paulsen, R., Bähner, M., Bentrop, J., Schillo, M., Schulz, S., Huber, A. (2001). The molecular design of a visual cascade: assembly of the Drosophila phototransduction pathway into a supramolecular signaling complex. In: Vision: the approach of biophysics and neurosciences (Musio, C., ed.), pp. 60–73. World Scientific, Singapore. 18. Montell, C. (1998). TRP trapped in fly signaling web. Curr. Opin. Neurobiol. 8, 389–397. 19. Huber, A. (2001). Scaffolding proteins organize multimolecular protein complexes for sensory signal transduction. Eur. J. Neurosci. 14, 769–776. 20. Ranganathan, R., Ross, E. M. (1997). PDZ domain proteins: scaffolds for signaling complexes. Curr. Biol. 7, R770–R773.

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21. Zuker, C. S., Cowman, A. F., Rubin, G. M. (1985). Isolation and structure of a rhodopsin gene from D. melanogaster. Cell 40, 851–858. 22. O’Tousa, J. E., Baehr, W., Martin, R. L., Hirsh, J., Pak, W. L., Applebury, M. L. (1985). The Drosophila ninaE gene encodes an opsin. Cell 40, 839–850. 23. Gehring, W. J. (2002). The genetic control of eye development and its implications for the evolution of the various eye-types. Int. J. Dev. Biol. 46, 65–73. 24. Nakagawa, M., Iwasa, T., Kikkawa, S., Tsuda, M., Ebrey, T. G. (1999). How vertebrate and invertebrate visual pigments differ in their mechanism of photoactivation. Proc. Natl. Acad. Sci. U. S. A. 96, 6189–6192. 25. Bentrop, J., Paulsen, R. (2003). Invertebrate rhodopsins. In: Photoreceptors and light signalling (Batschauer, A., ed.), pp. 40–76. Royal Society of Chemistry, Cambridge, UK. 26. Chyb, S., Raghu, P., Hardie, R. C. (1999). Polyunsaturated fatty acids activate the Drosophila light-sensitive channels TRP and TRPL. Nature 397, 255–259. 27. Pak, W. L., Grossfield, J., Arnold, K. S. (1970). Mutants of the visual pathway of Drosophila melanogaster. Nature 227, 518–520. 28. Pak, W. L. (1979). Study of photoreceptor function using Drosophila mutants. In: Neurogenetics: genetic approaches to the nervous system (Breakefield, X., ed.), pp. 67–99. Elsevier, Amsterdam. 29. Bloomquist, B. T., Shortridge, R. D., Schneuwly, S., Perdew, M., Montell, C., Steller, H., Rubin, G., Pak, W. L. (1988). Isolation of a putative phospholipase C gene of Drosophila, norpA, and its role in phototransduction. Cell 54, 723–733. 30. Shieh, B. H., Niemeyer, B. (1995). A novel protein encoded by the InaD gene regulates recovery of visual transduction in Drosophila. Neuron 14, 201–210. 31. Tsunoda, S., Sierralta, J., Sun, Y., Bodner, R., Suzuki, E., Becker, A., Socolich, M., Zuker, C. S. (1997). A multivalent PDZ-domain protein assembles signalling complexes in a G-proteincoupled cascade. Nature 388, 243–249. 32. Shieh, B. H., Zhu, M. Y. (1996). Regulation of the TRP Ca2+ channel by INAD in Drosophila photoreceptors. Neuron 16, 991–998. 33. Huber, A., Sander, P., Gobert, A., Bahner, M., Hermann, R., Paulsen, R. (1996). The transient receptor potential protein (Trp), a putative store-operated Ca2+ channel essential for phosphoinositide-mediated photoreception, forms a signaling complex with NorpA, InaC and InaD. EMBO J. 15, 7036–7045. 34. Huber, A., Sander, P., Paulsen, R. (1996). Phosphorylation of the InaD gene product, a photoreceptor membrane protein required for recovery of visual excitation. J. Biol. Chem. 271, 11710–11717. 35. Chevesich, J., Kreuz, A. J., Montell, C. (1997). Requirement for the PDZ domain protein, INAD, for localization of the TRP store-operated channel to a signaling complex. Neuron 18, 95–105. 36. Wes, P. D., Xu, X. Z., Li, H. S., Chien, F., Doberstein, S. K., Montell, C. (1999). Termination of phototransduction requires binding of the NINAC myosin III and the PDZ protein INAD. Nat. Neurosci. 2, 447–453. 37. Xu, X. Z., Choudhury, A., Li, X., Montell, C. (1998). Coordination of an array of signaling proteins through homo- and heteromeric interactions between PDZ domains and target proteins. J. Cell Biol. 142, 545–555. 38. Doyle, D. A., Lee, A., Lewis, J., Kim, E., Sheng, M., MacKinnon, R. (1996). Crystal structures of a complexed and peptide-free membrane protein-binding domain: molecular basis of peptide recognition by PDZ. Cell 85, 1067–1076. 39. Morais-Cabral, J. H., Petosa, C., Sutcliffe, M. J., Raza, S., Byron, O., Poy, F., Marfatia, S. M., Chishti, A. H., Liddington, R. C. (1996). Crystal structure of a PDZ domain. Nature 382, 649–652.

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Huber and Meyer

40. Hung, A. Y., Sheng, M. (2002). PDZ domains: structural modules for protein complex assembly. J. Biol. Chem. 277, 5699–5702. 41. Harris, B. Z., Lim, W. A. (2001). Mechanism and role of PDZ domains in signaling complex assembly. J. Cell Sci. 114, 3219–3231. 42. Songyang, Z., Fanning, A. S., Fu, C., Xu, J., Marfatia, S. M., Chishti, A. H., Crompton, A., Chan, A. C., Anderson, J. M., Cantley, L. C. (1997). Recognition of unique carboxyl-terminal motifs by distinct PDZ domains. Science 275, 73–77. 43. Stricker, N. L., Christopherson, K. S., Yi, B. A., Schatz, P. J., Raab, R. W., Dawes, G., Bassett, D. E., Jr., Bredt, D. S., Li, M. (1997). PDZ domain of neuronal nitric oxide synthase recognizes novel C-terminal peptide sequences. Nat. Biotechnol. 15, 336–342. 44. Maximov, A., Sudhof, T. C., Bezprozvanny, I. (1999). Association of neuronal calcium channels with modular adaptor proteins. J. Biol. Chem. 274, 24453–24456. 45. Kimple, M. E., Siderovski, D. P., Sondek, J. (2001). Functional relevance of the disulfide-linked complex of the N-terminal PDZ domain of InaD with NorpA. EMBO J. 20, 4414–4422. 46. Adamski, F. M., Zhu, M. Y., Bahiraei, F., Shieh, B. H. (1998). Interaction of eye protein kinase C and INAD in Drosophila. Localization of binding domains and electrophysiological characterization of a loss of association in transgenic flies. J. Biol. Chem. 273, 17713– 17719. 47. van Huizen, R., Miller, K., Chen, D. M., Li, Y., Lai, Z. C., Raab, R. W., Stark, W. S., Shortridge, R. D., Li, M. (1998). Two distantly positioned PDZ domains mediate multivalent INAD- phospholipase C interactions essential for G protein-coupled signaling. EMBO J. 17, 2285–2297. 48. Georgiev, P., Garcia-Murillas, I., Ulahannan, D., Hardie, R. C., Raghu, P. (2005). Functional INAD complexes are required to mediate degeneration in photoreceptors of the Drosophila rdgA mutant. J. Cell Sci. 118, 1373–1384. 49. Li, H. S., Montell, C. (2000). TRP and the PDZ protein, INAD, form the core complex required for retention of the signalplex in Drosophila photoreceptor cells. J. Cell Biol. 150, 1411–1421. 50. Tsunoda, S., Sun, Y., Suzuki, E., Zuker, C. (2001). Independent anchoring and assembly mechanisms of INAD signaling complexes in Drosophila photoreceptors. J. Neurosci. 21, 150–158. 51. Chorna-Ornan, I., Tzarfaty, V., Ankri-Eliahoo, G., Joel-Almagor, T., Meyer, N. E., Huber, A., Payre, F., Minke, B. (2005). Light-regulated interaction of Dmoesin with TRP and TRPL channels is required for maintenance of photoreceptors. J. Cell Biol. 171, 143–152. 52. Schulz, S., Huber, A., Schwab, K., Paulsen, R. (1999). A novel Gγ isolated from Drosophila constitutes a visual G protein γ subunit of the fly compound eye. J. Biol. Chem. 274, 37605–37610. 53. Yarfitz, S. L., Running, D. J., Froelick, G., Colley, N. J., Hurley, J. B. (1994). In situ assay of light-stimulated G protein activity in Drosophila photoreceptor G protein beta mutants. J. Biol. Chem. 269, 30340–30344. 54. Kosloff, M., Elia, N., Joel-Almagor, T., Timberg, R., Zars, T. D., Hyde, D. R., Minke, B., Selinger, Z. (2003). Regulation of light-dependent Gq alpha translocation and morphological changes in fly photoreceptors. EMBO J. 22, 459–468. 55. Cronin, M. A., Diao, F., Tsunoda, S. (2004). Light-dependent subcellular translocation of Gqalpha in Drosophila photoreceptors is facilitated by the photoreceptor-specific myosin III NINAC. J. Cell Sci. 117, 4797–4806. 56. Lee, S. J., Xu, H., Kang, L. W., Amzel, L. M., Montell, C. (2003). Light adaptation through phosphoinositide-regulated translocation of Drosophila visual arrestin. Neuron 39, 121–132.

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57. Bähner, M., Frechter, S., Da Silva, N., Minke, B., Paulsen, R., Huber, A. (2002). Lightregulated subcellular translocation of Drosophila TRPL channels induces long-term adaptation and modifies the light-induced current. Neuron 34, 83–93. 58. Shieh, B. H., Zhu, M. Y., Lee, J. K., Kelly, I. M., Bahiraei, F. (1997). Association of INAD with NORPA is essential for controlled activation and deactivation of Drosophila phototransduction in vivo. Proc. Natl. Acad. Sci. U. S. A. 94, 12682–12687. 59. Henderson, S. R., Reuss, H., Hardie, R. C. (2000). Single photon responses in Drosophila photoreceptors and their regulation by Ca2+. J. Physiol. (Lond.) 524, 179–194. 60. Huber, A., Sander, P., Bahner, M., Paulsen, R. (1998). The TRP Ca2+ channel assembled in a signaling complex by the PDZ domain protein INAD is phosphorylated through the interaction with protein kinase C (ePKC). FEBS Lett. 425, 317–322. 61. Liu, M., Parker, L. L., Wadzinski, B. E., Shieh, B. H. (2000). Reversible phosphorylation of the signal transduction complex in Drosophila photoreceptors. J. Biol. Chem. 275, 12194–12199. 62. Popescu, D. C., Ham, A. J., Shieh, B. H. (2006). Scaffolding protein INAD regulates deactivation of vision by promoting phosphorylation of transient receptor potential by eye protein kinase C in Drosophila. J. Neurosci. 26, 8570–8577. 63. Bähner, M., Sander, P., Paulsen, R., Huber, A. (2000). The visual G protein of fly photoreceptors interacts with the PDZ domain assembled INAD signaling complex via direct binding of activated Gαq to phospholipase Cβ. J. Biol. Chem. 275, 2901–2904. 64. Scott, K., Becker, A., Sun, Y., Hardy, R., Zuker, C. (1995). Gq alpha protein function in vivo: genetic dissection of its role in photoreceptor cell physiology. Neuron 15, 919–927. 65. Scott, K., Zuker, C. S. (1998). Assembly of the Drosophila phototransduction cascade into a signalling complex shapes elementary responses. Nature 395, 805–808. 66. Hardie, R. C., Martin, F., Cochrane, G. W., Juusola, M., Georgiev, P., Raghu, P. (2002). Molecular basis of amplification in Drosophila phototransduction: Roles for G protein, phospholipase C, and diacylglycerol kinase. Neuron 36, 689–701. 67. Baylor, D. A., Matthews, G., Yau, K. W. (1980). Two components of electrical dark noise in toad retinal rod outer segments. J. Physiol. 309, 591–621. 68. Elia, N., Frechter, S., Gedi, Y., Minke, B., Selinger, Z. (2005). Excess of Gβe over Gqαe in vivo prevents dark, spontaneous activity of Drosophila photoreceptors. J. Cell Biol. 171, 517–526. 69. Cook, B., Bar-Yaacov, M., Cohen, B., Goldstein, R. E., Paroush, Z., Selinger, Z., Minke, B. (2000). Phospholipase C and termination of G-protein-mediated signalling in vivo. Nat. Cell Biol. 2, 296–301. 70. Molday, R. S. (1996). Calmodulin regulation of cyclic-nucleotide-gated channels. Curr. Opin. Neurobiol. 6, 445–452. 71. Körschen, H. G., Beyermann, M., Muller, F., Heck, M., Vantler, M., Koch, K. W., Kellner, R., Wolfrum, U., Bode, C., Hofmann, K. P., Kaupp, U. B. (1999). Interaction of glutamicacid-rich proteins with the cGMP signalling pathway in rod photoreceptors. Nature 400, 761–766. 72. Poetsch, A., Molday, L. L., Molday, R. S. (2001). The cGMP-gated channel and related glutamic acid-rich proteins interact with peripherin-2 at the rim region of rod photoreceptor disc membranes. J. Biol. Chem. 276, 48009–48016. 73. Molday, R. S., Molday, L. L. (1998). Molecular properties of the cGMP-gated channel of rod photoreceptors. Vision Res. 38, 1315–1323. 74. Bauer, P. J., Drechsler, M. (1992). Association of cyclic GMP-gated channels and Na(+)Ca(2+)-K+ exchangers in bovine retinal rod outer segment plasma membranes. J. Physiol. 451, 109–131.

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Huber and Meyer

75. Schwarzer, A., Kim, T. S., Hagen, V., Molday, R. S., Bauer, P. J. (1997). The Na/Ca-K exchanger of rod photoreceptor exists as dimer in the plasma membrane. Biochemistry. 36, 13667–13676. 76. Schwarzer, A., Schauf, H., Bauer, P. J. (2000). Binding of the cGMP-gated channel to the Na/Ca-K exchanger in rod photoreceptors. J. Biol. Chem. 275, 13448–13454. 77. Boesze-Battaglia, K., Lamba, O. P., Napoli, A. A., Jr., Sinha, S., Guo, Y. (1998). Fusion between retinal rod outer segment membranes and model membranes: a role for photoreceptor peripherin/rds. Biochemistry 37, 9477–9487.

13 Visual Signal Processing in the Inner Retina Botir T. Sagdullaev, Tomomi Ichinose, Erika D. Eggers, and Peter D. Lukasiewicz CONTENTS Introduction Visual Information Is First Processed in the OPL Bipolar Cells Form Parallel Pathways and Provide Excitatory Input to the IPL Functional Stratification of the IPL Synaptic Mechanisms Shape Excitatory Signals in the IPL Presynaptic Inhibition Conclusions References

INTRODUCTION The inner plexiform layer (IPL) is the second synaptic layer of the retina (Fig. 1) and the final stage for processing visual information before it leaves the eye. Visual signals from rod and cone photoreceptors are first processed in the outer plexiform layer (OPL; Fig. 1), where horizontal cells modulate their signaling to bipolar cells. Bipolar cells transmit these signals to the IPL, where amacrine cells shape bipolar cell signaling to ganglion cells. Bipolar cells form parallel sensory pathways that transmit information about specific aspects of the visual world to the IPL. Synaptic interactions between bipolar, amacrine, and ganglion cells in the IPL result in complex processing of the visual signals that are crucial for the detection of motion, contrast, color, and dim illumination. New evidence is starting to reveal how synaptic interactions in the IPL shape these parallel signals. The output of the IPL is then transmitted to distinct ganglion cell types that convey visual signals to different brain regions. This chapter reviews the current knowledge of how the IPL shapes visual signaling.

From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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Fig. 1. A cross-sectional view of retinal layers and the main vertical and lateral signaling pathways. The retina is organized into three cellular layers, rod (R) and cone (C) photoreceptor somas are located in the outer nuclear layer (ONL); horizontal cell (HC), bipolar cell (BC), and amacrine cell (AC) somas are located in the inner nuclear layer (INL); ganglion cell (GC) somas are located in the ganglion cell layer (GCL). Synaptic connections between photoreceptors, bipolar cells, and horizontal cells are made in the outer plexiform layer (OPL). Synaptic connections between bipolar cells, amacrine cells, and ganglion cells are made in the inner plexiform layer (IPL). Vertical signaling pathways are composed of photoreceptors, bipolar cells, and ganglion cells. Two lateral pathways, comprised of horizontal cells in the OPL and amacrine cells in the IPL, modulate the flow of information along the vertical pathway.

VISUAL INFORMATION IS FIRST PROCESSED IN THE OPL Photoreceptors transduce light into electrical signals, which are processed by retinal interneurons before being sent to the brain. In the first synaptic layer, horizontal cells make synaptic contacts with photoreceptor terminals (Fig. 1). One outcome of this interaction is the establishment of an antagonistic center/surround receptive field organization, which can be revealed by stimulating the retina with light spots of different sizes. Illumination of the receptive field center with a small spot of light alters the activity of photoreceptors, which integrate inputs over small areas. Illumination of the larger receptive field surround alters the activity of horizontal cells, which feed back to the photoreceptor, antagonizing the effects of center illumination [1]. The antagonistic center/surround organization of receptive field is a fundamental feature of bipolar and ganglion cells [2, 3]. In the retina, this receptive field organization contributes to contrast perception, edge detection, and color processing [4, 5]. Additional center-surround signal processing also occurs in the IPL, as described next. BIPOLAR CELLS FORM PARALLEL PATHWAYS AND PROVIDE EXCITATORY INPUT TO THE IPL At the OPL, the processed photoreceptor output is relayed to the IPL by a variety of bipolar cell types that form parallel signaling pathways (reviewed in [6, 7]). Excitatory inputs to the IPL originate from the two main functional classes, ON and OFF bipolar cells. ON bipolar cells depolarize in response to increasing illumination within their receptive field centers, and OFF bipolar cells depolarize in response to decreasing illumination within their receptive field centers. The distinct responses of

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Fig. 2. Bipolar cells form parallel retinal signaling pathways. Vertical view drawings of neurobiotin stained bipolar cells from rat retina illustrate distinct morphological subtypes. A similar diversity of bipolar cells exists in retinas of other mammalian and cold-blooded vertebrates. The axon terminals stratify at different depths of the inner plexiform layer (IPL). ON bipolar cells have axons that terminate in the inner half of the IPL (ON sublamina), and OFF bipolar cells have axons that terminate in the outer half of the IPL (OFF sublamina). Rod bipolar cell (RB) dendrites exclusively contact rods. The remaining cone bipolar cells (CB) only contact cones. GCL ganglion cell layer, INL inner nuclear layer, OPL outer plexiform layer (Used with permission of the American Physiological Society, from [47].)

ON and OFF bipolar cells are determined by separate classes of glutamate receptors present on their dendrites. Glutamate is continuously released by photoreceptors in the dark and depolarizes OFF bipolar cells by activating α-amino-3-hydroxy-5-methyl-4isoxazol-propionic acid (AMPA) and kainate types of ionotropic glutamate receptors on their dendritic processes [8, 9]. ON bipolar cells, by contrast, are hyperpolarized by glutamate that activates dendritic metabotropic mGluR6 (metabotropic glutamate receptor 6) receptors [10, 11]. In the light, when glutamate release is decreased, ON bipolar cells depolarize, and OFF bipolar cells hyperpolarize. These broad ON and OFF bipolar cell classes are divided further into several subtypes of ON and OFF bipolar cells (reviewed in [6]). In mammalian retina, at least nine morphological classes of ON and OFF bipolar cells receive input from cones, and one class receives input exclusively from rods (Fig. 2). A similar variety of ON and OFF bipolar cells exists in salamander; however, these bipolar cells receive mixed inputs and are either cone or rod dominant [12]. The morphological subtypes of ON and OFF bipolar cells have unique functions. In mammals, ON rod bipolar cells contact rods exclusively and transmit rod signals to the IPL. The remaining bipolar cell types relay different cone signals to the IPL. Some types of cone bipolar cells relay color information between cones and ganglion cells. For instance, midget bipolar cells comprise the red-green signaling pathway and transmit red and green spectral information. The blue cone bipolar cells receive input exclusively from blue cones and comprise the blue pathway. Another cone bipolar cell class, called diffuse bipolar cells, transmits luminosity information without encoding spectral information (“color blind”). The electrical responses of separate bipolar cell classes to photoreceptor input are distinct, consistent with the notion that bipolar cells separate the visual input into parallel signals. How is the photoreceptor input separated into different signals? Distinct gluta-

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Fig. 3. Photoreceptor inputs are differentially filtered by bipolar cell glutamate receptors. A Schematic showing b2 and b3 bipolar cells contacting a single cone. B Depolarization of the cone elicited responses in b2 and b3 bipolar cells that were simultaneously recorded (upper panel). Scaled versions of the simultaneously recorded responses (lower panel) illustrate the difference in response time course, which is attributable to differential filtering by the b2 and b3 bipolar cell synapses. (Panel B is reprinted from [8], with permission from Elsevier.)

mate receptor subtypes differentially filter photoreceptor input in distinct morphological bipolar cell types to transmit different features of the visual input, as illustrated in Fig. 3 [8–10]. This filtering causes some bipolar cells to respond transiently and other bipolar cells to respond in a sustained fashion, enabling separate bipolar cells to extract distinct temporal and spatial features of the visual input. Further separation of color and luminosity information also occurs by distinct bipolar cells [13]. These morphologically and functionally distinct bipolar cell types transmit different representations of the visual scene to the IPL for additional processing [14]. FUNCTIONAL STRATIFICATION OF THE IPL Bipolar cells convey different representations of photoreceptor input to the IPL by slow, graded potentials. When the ON and OFF classes of bipolar cells are depolarized by increments or decrements of illumination, respectively, they release glutamate, which excites postsynaptic ganglion cells and amacrine cells. The excitatory output of each bipolar cell class is shaped by amacrine cell synaptic inputs to bipolar cell terminals, as described in detail, the section on Presynaptic Inhibition. Additional signal processing, attributable to amacrine and ganglion cell interactions, also occurs in the IPL, before visual information is sent to the higher visual centers. ON and OFF Response Stratification The axon terminals of ON and OFF bipolar cells stratify in two distinct sublaminae of the IPL, where they contact the dendrites of ON and OFF ganglion and amacrine cells. These two strata are where synaptic connections are made by the pathways that encode

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visual responses to increments and decrements in intensity across the visual scene (Fig. 2). The outer half of the IPL encodes responses to decrements (the OFF sublaminae), and the inner half encodes responses to increments in illumination (the ON sublaminae) [15, 16]. In the ON sublaminae, synaptic contacts are made between bipolar and ganglion cells that are excited by increments of illumination of their receptive field centers. In the OFF sublaminae, synaptic connections are made between bipolar and ganglion cells that are excited by decrements of illumination of their receptive field centers. The ON and OFF sublaminae are further divided into ten morphological strata (see Fig. 2) that are functionally distinct and encode different representations of the visual scene [14]. Sustained and Transient Response Stratification Some retinal neurons generate brisk, transient responses to continuous visual stimulation, while others maintain their activity for the duration of the stimulus. These transient and sustained cell classes are believed to play crucial roles in encoding the temporal and spatial features of the visual world and represent additional parallel sensory channels in the retina. A major functional subdivision of the ON and OFF sublaminae is the sustained and transient response stratification that occurs in each sublaminae, suggesting that there are transient and sustained responding subtypes of the major ON and OFF classes of retinal neurons. Sustained and transient strata were first described in turtle retina [17] and subsequently confirmed in other species [10, 12, 14]. Sustained and transient bipolar cells provide the inputs to these IPL strata [10]. Transient bipolar cell axon terminals contact transient amacrine and ganglion cell processes in the mid-IPL, and sustained bipolar cells axon terminals contact the sustained responding amacrine and ganglion cells near the inner and outer margins of the IPL [10, 12, 18]. SYNAPTIC MECHANISMS SHAPE EXCITATORY SIGNALS IN THE IPL The functional stratification of the IPL suggests that distinct bipolar cells generate sustained and transient responses. Wunk and Werblin [19] suggest that the separation of sustained and transient visual signals occurs in the IPL, where synaptic interactions generate sustained and transient bipolar cell outputs. However, subsequent work suggests that sustained and transient visual signals are generated in the OPL at the dendrites of different bipolar cell subtypes, attributable to the filtering of distinct glutamate receptors [9, 10]. Although the main signal separation occurs in the OPL, additional refinement of sustained and transient signals takes place in the IPL. Glutamate release from bipolar cells is truncated by inhibitory amacrine cell input to shape the time course of transient excitatory responses in ganglion cells [20–22]. As noted in section, Synaptic Mechanisms Shape Excitatory Signals in the IPL, transient ganglion cell responses are also shaped by other IPL synaptic mechanisms (see Fig. 4B), such as glutamate uptake by transporters [23] and desensitizing glutamate receptors [24]. Glutamate Release Is Tonic and Graded Since bipolar cells do not use action potentials, but use slow graded depolarizations to signal, glutamate release is graded and tonic [25, 26]. The release machinery in bipolar cells is optimized for sustained glutamate release. The L-type calcium channels, present

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Fig. 4. A A bipolar cell terminal illustrating the processes that contribute to tonic glutamate release. Tonic release is attributed to sustained, graded depolarizations, prolonged calcium influx through L-type calcium channels, and a large pool of ribbon-associated vesicles. B A bipolar cell terminal illustrating the synaptic mechanisms that shape excitatory signaling to ganglion cells. Transporters limit excitatory signaling by removing glutamate from the synapse. Desensitizing postsynaptic glutamate receptors (X) limit responses to sustained glutamate release. Glutamate release may be reduced by the activation of presynaptic metabotropic glutamate receptors (mGluR) or the activation of presynaptic γ-aminobutyric acid A (GABAA) and GABAC receptors.

in bipolar terminals, open in response to graded depolarization and mediate a sustained calcium influx that elicits tonic glutamate release (Fig. 4A). Specialized structures called synaptic ribbons are located at release zones and contain large numbers of tethered, glutamate-filled vesicles (Fig. 4A) that mediate sustained signaling (for review, see [27]). While tonic glutamate release is tailored for graded signaling to ganglion cells, there are several challenges that this signaling poses. How is the sustained signal rapidly terminated? Do postsynaptic glutamate receptors desensitize to tonic glutamate release? These mechanisms are considered in more detail. Transporters Terminate Excitatory Signaling to Ganglion Cells In most parts of the central nervous system (CNS), synaptic responses are terminated by either the chemical degradation of transmitter or, in case of glutamate, by its rapid diffusion from the synaptic cleft [28, 29]. However, at bipolar-to-ganglion cell synapses [23, 30], and other specialized synapses in the CNS [31–33], excitation is terminated by the active clearance of glutamate by transporters into surrounding neurons and glia. Glutamate transporters shape ganglion cell excitatory responses (Fig. 4B). Blockade of glutamate transporters in the IPL enhances and prolongs glutamate signaling to ganglion cells, indicating that transporters limit the amplitude and time course of ganglion cell excitation [23]. Postsynaptic Glutamate Receptor Properties Shape Ganglion Cell Excitation Glutamate receptors on ganglion cells are exposed to sustained, elevated glutamate concentrations, attributable to sustained light-evoked release and slow clearance by

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transporters (compared to diffusion). Glutamate activates both AMPA and NMDA (N-methyl-D-aspartate) receptors on ganglion cells. AMPA receptors desensitize to sustained activation by glutamate (Fig. 4B), responding transiently to maintained stimuli [34]. When AMPA receptor desensitization is reduced in ganglion cells, either pharmacologically or by holding the cells at positive potentials, excitatory light responses are enhanced, suggesting that AMPA receptor desensitization shapes ganglion cell excitatory responses [23, 24]. Both AMPA receptor desensitization and glutamate uptake limit ganglion cell excitation, but they do so in distinct ways. Glutamate uptake limits the time course of excitation, and receptor desensitization limits the amplitude of the late phase of excitation [23]. Modulating Glutamate Release Shapes Excitatory Responses Several mechanisms that modulate glutamate release from bipolar cells control excitatory signaling to ganglion cells. These mechanisms include autoinhibitory mechanisms by which glutamate and protons that are released from exocytosed vesicles feed back to limit glutamate release [35–37]. Glutamate limits release by activating metabotropic glutamate receptors, and protons limit release by inhibiting calcium influx (Fig. 4B). Also, as described next, presynaptic inhibition by amacrine cells also limits the probability of glutamate release [20, 38]. Amacrine Cells Mediate Inhibition in the IPL While excitatory input to ganglion cells comes almost exclusively from bipolar cells, inhibitory signals in the IPL are mediated by amacrine cells. Amacrine cells are the most diverse class of retinal interneurons (Fig. 5). They are morphologically and functionally distinct, with different sets of neurotransmitters and receptors. Amacrine cells, like ganglion cells, are excited by bipolar cells. They mediate inhibition by releasing either γ-aminobutyric acid (GABA) or glycine onto their postsynaptic targets, which include ganglion cell dendrites, bipolar cell axon terminals, and other amacrine cells. Approximately 50% of amacrine cells are GABAergic, and 50% are glycinergic. Ganglion cells receive mainly amacrine cell input (Masland 2001), underscoring the importance of inhibition in shaping the output of the retina. Bipolar cell terminals are inhibited by presynaptic amacrine cell inputs, controlling glutamate release and excitatory signaling to ganglion cells. Amacrine cells also inhibit other amacrine cells, resulting in complex serial, inhibitory synaptic interactions [39–41]. There is also a population of excitatory amacrine cells called starburst amacrine cells, but their role in visual signal processing in adult animals remains unclear (reviewed in [42]) and is considered in the section, Directional-Selective Ganglion Cells. There are two fundamental, morphological classes of amacrine cells, narrow field and wide field, named for the extent of their processes (Fig. 5). Wide-field amacrine cells are composed of many functional and neurochemical (~15) subtypes [6]. However, they share the common attribute of signaling over relatively long distances, usually confined to specific IPL strata. The narrow-field amacrine cell class is also composed of many subtypes that mediate local signaling. Another trait of narrow-field amacrine cells is that they often signal between strata, mediating vertical interaction across different layers within the IPL. Thus, communication between strata that represent distinct functional channels is mediated by narrow-field amacrine cells [43].

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Fig. 5. The major types of amacrine cells in mammalian retina are shown. Amacrine cells comprise the most diverse class of retinal neurons. This illustrates amacrine cells that were morphologically identified in rabbit retina. Narrow-field amacrine cells are shown in the top row, and wide-field amacrine cells are shown in the lower two rows. (Reprinted from [73], with permission from Elsevier.)

PRESYNAPTIC INHIBITION Presynaptic inhibition of bipolar cell terminals by amacrine cells exists in two general forms and contributes to the temporal and spatial properties of visual processing. Local presynaptic inhibition, also called feedback inhibition, mediated by reciprocal synapses between bipolar and amacrine cells shapes the time course [20] and extent of glutamate release [22]. Lateral inhibition, mediated by long-distance presynaptic inhibitory signaling from amacrine cells, contributes to the antagonistic receptive field surround of ganglion cells [44–46]. Asymmetric Presynaptic Inhibition Both ON and OFF bipolar cells receive presynaptic input from amacrine cells, suggesting that signaling to ON and OFF ganglion cells is shaped by presynaptic inhibition. However, presynaptic inhibition differentially shapes ON and OFF pathway signaling in the IPL [22]. This was demonstrated by eliminating the main type of presynaptic inhibition to bipolar cells. When GABAC receptor-mediated presynaptic inhibition was eliminated, Sagdullaev and colleagues showed that ON but not OFF ganglion cell responses were greatly enhanced, suggesting that presynaptic inhibition was asymmetric (Fig. 6). Electrophysiological measurements of light- and electrically evoked excitation to ganglion cells showed that presynaptic inhibition affected the dynamic response ranges in ON ganglion cells by limiting glutamate release from ON but not OFF bipolar cells. Presynaptic inhibition of ON bipolar cells modulates the dynamic response range by limiting the extent of glutamate spillover and activation of perisynaptic NMDA receptors (Fig. 6). Spillover transmission occurs at some synapses, including ON ganglion cells, when glutamate diffuses from the release sites and activates perisynaptic receptors. Recent evidence suggests that spillover activation of NMDA receptors occurs only at ON ganglion cells (Fig. 7) [22]. When glutamate release is increased or glutamate uptake is blocked, glutamate concentrations are increased, and spillover is enhanced. Sagdullaev and colleagues found that manipulations that increased spillover enhanced the activation of NMDA receptors in ON but not OFF ganglion cells. These findings suggest another

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asymmetry is present in the IPL; NMDA receptors are located perisynaptically in ON and synaptically in OFF ganglion cell synapses (Fig. 6). Presynaptic Inhibition Is Filtered by GABA Receptor Properties GABAergic inhibition occurs at the terminals of all classes of bipolar cell. Pharmacological studies indicated that two types of ionotropic GABA receptors, GABAA and GABAC receptors, are present on bipolar terminals [47, 48]. Pharmacologically isolated GABAA and GABAC receptors mediate distinct inhibitory responses to applied GABA [48]. GABAA receptor-mediated responses rise and decay rapidly, while GABAC receptor-mediated responses rise and decay slowly (Fig. 7A). Do these distinct GABA receptor properties shape light-evoked inhibitory responses? We found that these distinct GABA receptors temporally filter synaptic input to bipolar cells [41]. Recordings of light-evoked inhibitory postsynaptic currents (L-IPSCs) from rod bipolar cells demonstrate that slowly responding GABAC receptors prolong the response decay (Fig. 7B). By contrast, the rapidly activating and decaying GABAA receptors (Fig. 7B) determine the rise time and peak amplitude of the response [49]. GABA application experiments indicated that diverse classes of bipolar cells have different proportions of GABA A and GABAC receptors [47, 48], suggesting that light-evoked inhibition is differentially filtered at different classes of bipolar cells. Eggers and colleagues [50] showed that the L-IPSC time course varied in different bipolar cell classes, depending on the relative contributions of GABAA and GABAC receptors. These findings suggest that GABAergic L-IPSCs are temporally filtered by distinct receptors; GABAC receptor-mediated L-IPSCs are prolonged, and GABAA receptor-mediated L-IPSCs are brief. GABAergic L-IPSCs recorded in rod bipolar cells decay slowly because they are dominated by GABAC receptors, while OFF cone bipolar cell L-IPSCs decayed rapidly, reflecting a larger GABAA receptor contribution. These observations demonstrate that different GABA receptor complements differentially tune inhibition for specific bipolar cell types. These two forms of presynaptic inhibition limit bipolar cell outputs in distinct ways. Slow GABAC receptors limit the extent of glutamate release and the duration excitatory responses in amacrine and ganglion cells. Fast GABAA receptors, by contrast, limit the initial glutamate release and the initial postsynaptic excitatory responses. Different complements of GABAA and GABAC receptors also appear to match the time course of inhibition with the time course of photoreceptor input to different bipolar cells. Rod bipolar cells receive prolonged excitatory input from rod photoreceptors and prolonged presynaptic inhibition mediated by GABAC receptors. OFF cone bipolar cells receive brisk excitatory input from cone photoreceptors and fast presynaptic inhibition mediated by GABAA receptors. Presynaptic Inhibition May Be Shaped by Transmitter Release Differences It is not known whether GABAA and GABAC receptors receive input from similar or distinct presynaptic amacrine cell inputs. To address this issue, Eggers and colleagues [49] used deconvolution analysis [51] to estimate the GABA release time courses associated with GABAA and GABAC receptor-mediated light-evoked currents. They found that the apparent release time courses for inputs to GABAA and GABAC receptor-containing synapses were distinct. Although this finding is consistent with

Fig. 6. Asymmetric presynaptic inhibition differentially affects ON and OFF pathway signaling. Raster plots (upper traces) and peristimulus time histograms (PSTHs; middle) illustrating spontaneous and light-evoked firing in WT (Ai) and GABACR null (Null) (Aii) ON-center ganglion cells (GCs); and WT (Bi) and Null (Bii) OFF GCs. The lower traces in (A) and (B) indicate the duration of the stimuli, a bright, centered spot for ON GCs and a dark, centered spot for OFF GCs, presented on an adapting background. Spontaneous and light-evoked firing rates were significantly increased only in ON GCs in Null mice compared to WT mice. C The asymmetric presynaptic inhibition of glutamate release from bipolar cells (BCs) and spillover activation of postsynaptic NMDA (N-methyl-D-aspartate) receptors (NMDARs)on GCs. Ci GABAergic feedback from amacrine cells (ACs) limits glutamate release from ON bipolar cells and limits spillover activation of perisynaptic (NMDARs) on ON GCs dendrites. GABACR-mediated negative feedback confines synaptic transmission and extends the dynamic response range of ON GCs. Cii When GABACR-mediated inhibition is eliminated, the modulation of the excitatory transmission is disrupted and glutamate release is enhanced. D In the OFF pathway, the activation of synaptically localized AMPARs and NMDARs on OFF GC dendrites is not limited by GABACR-mediated feedback to OFF BCs. The output gain of OFF GCs is high because their excitatory inputs are not appreciably modulated by presynaptic inhibition. For simplicity, only the inhibitory feedback component of a reciprocal synapse between a BC and an AC is shown. GABA γ-aminobutyric acid RGC retinal ganglion cell (Reprinted from [22], with permission from Elsevier.)

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Fig. 7. γ-Aminobutyric acid- (GABA-) and light-evoked inhibition at bipolar cell axon terminals is shaped by GABA receptor properties. A Whole-cell recording of currents in response to GABA puffed onto bipolar cell axon terminals. B GABAA and GABAC receptor-mediated currents have distinct time courses. GABAA receptor-mediated responses were isolated using the GABAC receptor antagonist 1,2,5,6-Tetrahydropyridin-4-yl methylphosphinic acid (TPMPA) and GABAC receptor-mediated responses were isolated using the GABAA receptor antagonist bicuculline. A cocktail of glycine and glutamate receptor antagonists was used to isolate the GABA receptor response components. C Light stimulation and recording procedure for measuring GABAergic L-IPSCs in bipolar cells. D. Pharmacologically isolated GABAA- and GABAC-receptor mediated light-evoked inhibitory postsynaptic currents (L-IPSCs) recorded from a bipolar cell, scaled to the same peak amplitude to compare response kinetics. Similar to the puff-evoked responses, the GABAA receptor-mediated L-IPSC exhibited fast rise and decay times, while the GABAC receptor-mediated L-IPSC exhibited slow rise and decay times. GABA-mediated L-IPSCs were isolated by pharmacologically blocking glycine receptors.

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distinct inputs, the release time course sensed by GABAC receptors may be distorted by the saturation and slow deactivation of these receptors, obscuring a common input. Inspection of the apparent GABAC receptor release function reveals a prolonged tail, not observed with GABAA receptors, which is consistent with the spillover activation of GABAC receptors. Because GABAC receptors are more sensitive to GABA than GABAA receptors, they may be activated by low concentrations of GABA that spill over from neighboring release sites. When GABA spillover is enhanced, by either reducing uptake or enhancing release, the GABAC, but not the GABAA, receptor-mediated component of light-evoked inhibition is selectively increased [52]. GABA spillover leads to stronger GABACR-mediated presynaptic inhibition, larger reductions in glutamate release, and reduced ganglion cell excitation [52]. Taken together, these data suggest that, in addition to slow receptor kinetics of GABACRs, spillover contributes to the prolonged GABACRmediated responses. Glycine, the Other Inhibitory Transmitter About one half of all amacrine cells are glycinergic, and there are eight types of glycinergic amacrine cells [53]. Anatomical evidence suggests that rod and OFF cone bipolar cells receive input from glycinergic amacrine cells [54, 55]. Physiological recordings of spontaneous [56] and light-evoked [74] glycinergic inhibitory currents have shown that presynaptic inhibition was most prominent in OFF cone bipolar cells and was also observed in rod bipolar cells, but not in ON cone bipolar cells. Slow rod signals are transmitted from rod bipolar cells to OFF bipolar cell terminals by AII (or rod) amacrine cells. This light-evoked presynaptic inhibition is slow, matches the time course of rod signaling, and is mediated by glycine [74]. There is an apparent mismatch between the slow L-IPSC time course and fast kinetics of the glycine receptors [56]. This discrepancy can be reconciled when one considers that the time course of glycine release from AII amacrine cells is likely to be sustained, similar to its sustained light-evoked depolarization [57, 58]. Thus, the time course of glycinergic inhibition is matched to the slow rod signal time course, but the time course is determined by transmitter release and not receptor properties. Lateral Versus Vertical Inhibitory Pathways in the IPL: The Story of Two Inhibitory Neurotransmitters Why does the IPL utilize two inhibitory transmitters, GABA and glycine? Although both of these transmitters gate chloride channels, these two transmitters are utilized by distinct signaling pathways. In mammalian retina, GABAergic amacrine cells have wide-field processes and are confined to the same laminae and signal laterally [6]. By contrast, the processes of glycinergic amacrine cells have narrow extents, but are typically multistratified and signal vertically between different laminae [6, 53]. GABAergic lateral inhibition plays a critical role in the processing of spatial information. Wide-field GABAergic amacrine cells contribute to the receptive field surround of ganglion cells [44, 45]. Cook and McReynolds [44] suggested that amacrine cell surround input mediates a finer spatial filtering that occurs after the coarser spatial filtering in the OPL, attributed to horizontal cell surround signaling. Wide-field GABAergic amacrine

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cells synapse onto bipolar cell terminals and ganglion cell dendrites. Thus, there are two IPL surround signaling pathways: direct GABAergic inhibition of ganglion cells [44] and presynaptic GABAergic inhibition that limits bipolar cell signaling to ganglion cells [45, 46]. It remains to be seen whether similar or different classes of amacrine cells mediate pre- and postsynaptic wide-field inhibition. Glycinergic amacrine cells have narrow-field processes that extend vertically across the IPL and mediate signaling between different sublamina [14, 43]. The best-described glycinergic amacrine cell is the AII or rod amacrine cell. Rod bipolar cells do not directly contact ganglion cells. Instead, they signal through an AII amacrine cell intermediate that contacts ON and OFF cone bipolar cells, which in turn signal ganglion cells. The AII amacrine splits the ON rod bipolar cell input into an inhibitory glycinergic output to OFF cone bipolar cell terminals [59] and excitatory, electrical output to ON cone bipolar cell terminals, mediated by gap junctional connections [60]. In contrast to wide-field GABAergic amacrine cells that signal laterally within IPL strata, narrow-field glycinergic cells signal vertically between IPL strata. Parallel Ganglion Cell Output Pathways Parallel bipolar cell-signaling pathways relay different aspects of the visual signal to 10–15 morphological types of ganglion cells [6, 61]. As noted, amacrine cells modulate the information transferred between bipolar and ganglion cells. This information is then sent by distinct ganglion cells that comprise parallel signal pathways to different parts of the brain. Like bipolar cells, ganglion cells are divided into two major ON and OFF classes, which convey information about light increments and decrements in their receptive field’s center, respectively, and both classes possess antagonistic surrounds [2]. ON and OFF types can be further divided into X and Y cells that summate inputs linearly and nonlinearly, respectively [62]. They respond to illumination in either a sustained (X) or transient (Y) fashion, similar to bipolar cells. In cat retina, these Y and X ganglion cell classes form distinct morphological types, α- and β-ganglion cells, respectively [63]. Transient bipolar cell terminals and Y ganglion cell dendrites costratify in the middle IPL strata, while sustained bipolar cell terminals and X ganglion cell dendrites costratify at the inner- and outermost strata of the IPL. The X and Y ganglion cells are similar to the parvo (P) and magno (M) classes of primate ganglion cells, which correspond to the morphological midget and parasol classes, respectively (reviewed in [64]). Parvo ganglion cells transmit visual information about form and color (what is it?), while magno ganglion cells transmit information about motion and spatial relationship (where is it?) to the lateral geniculate nucleus (reviewed in [65]). Ganglion Cells Encode Color Information Midget ganglion cells encode color information and relay it to the brain. Like the midget bipolar cells, midget ganglion cells comprise the red-green pathway. This redgreen color opponent pathway is most pronounced in the central retina and is attributed to the unique circuitry of the midget system. Their center response is determined by a strong single red or green cone input, while the surround is determined by weak mixed cone inputs, resulting in color opponency. A unique bistratified ganglion cell that is excited by blue light, via blue bipolar cells, makes contacts at the inner IPL, and that

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is inhibited by yellow light, via red and green signaling bipolar cells, that makes contacts at the outer IPL to process blue-yellow color opponency. Directional-Selective Ganglion Cells A subset of ganglion cells responds to the movement of images, but only in preferred directions. These ganglion cells are called directionally selective (D-S) cells and are a favorite of retinal electrophysiologists because they serve as model output cells that reveal the complexity of synaptic computation in the IPL. Although D-S ganglion cells were first described by Barlow and colleagues [66] over 40 years ago, we still do not know the synaptic interactions responsible for their behavior. What is known is that image motion in the preferred direction excites D-S cells, which are then inhibited after a time lag. By contrast, image motion in the null direction elicits a long-lasting inhibition that shunts the excitatory input, resulting in no response. One study suggested that this D-S computation occurs at the level of the ganglion cell dendrites, which integrate excitatory and inhibitory inputs [67]. However, other studies challenged this notion and suggested that the input neurons to the D-S cells are themselves directionally selective [68–70]. One point of agreement between most studies is that starburst amacrine cells are critical for D-S responses in ganglion cells. The genetic ablation of starburst cells, which provide both cholinergic and GABAergic input to D-S cells, results in the elimination D-S responses [71]. Thus, starburst amacrine cells are most likely critical players in the formation of D-S ganglion cell response, but precise roles of presynaptic and postsynaptic computations still need to be worked out. Intrinsically Photosensitive Ganglion Cells Recent work revealed that 1–3% of ganglion cells contain the photopigment melanopsin and can directly sense light [72]. These ganglion cells respond to light even when rod and cone inputs are disrupted, consistent with the notion that they are intrinsically photosensitive. Intrinsic light responses are long lasting and activated by moderateto-bright light intensities. This class of ganglion cell also receives input from highersensitivity rod and cone synaptic pathways, which elicit responses by these neurons at lower light intensities. These ganglion cells have extensive dendritic fields that completely tile the retina, allowing this system to respond to light impinging on any part of the retina. Intrinsically photosensitive ganglion cells project to areas of the brain that control pupillary reflexes and the light entrainment of circadian rhythms. CONCLUSIONS The IPL is a critical stage in visual processing. Parallel streams of information arrive in the IPL via distinct bipolar cell types. This information is then further shaped by synaptic interaction between inhibitory amacrine cells with the bipolar cell inputs and ganglion cell outputs of the IPL. The ability to detect the direction of movement and certain colors occurs because of processing in the IPL. Many questions about IPL processing still remain. The challenge of future research is to apply new techniques such as molecular genetic approaches of disrupting circuitry and powerful multielectrode recording techniques to gain additional insights into IPL function.

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REFERENCES 1. Baylor DA, Fuortes MGF, O’Bryan PM. Receptive fields of single cones in the retina of the turtle. J Physiol (Lond) 1971;214:265–294. 2. Kuffler SW. Discharge patterns and functional organization of mammalian retina. J Neurophys 1953;16:37–68. 3. Werblin FS, Dowling JE. Organization of the retina of mudpuppy, Necuturus maculosis. II. Intracellular recording. J Neurophys 1969;32:339–355. 4. Stell W, K, Lightfoot D, O. Color-specific interconnections of cones and horizontal cells in the retina of goldfish. J Comp Neurol 1975;159:473–502. 5. Werblin FS. The control of sensitivity in the retina. Sci Am 1973;228:70–79. 6. Wassle H. Parallel processing in the mammalian retina. Nat Rev Neurosci 2004;5(10): 747–757. 7. Masland RH. The fundamental plan of the retina. Nat Neurosci 2001;4(9):877–886. 8. Devries SH, Li W, Saszik S. Parallel processing in two transmitter microenvironments at the cone photoreceptor synapse. Neuron 2006;50(5):735–748. 9. DeVries SH. Bipolar cells use kainate and AMPA receptors to filter visual information into separate channels. Neuron 2000;28(3):847–856. 10. Awatramani GB, Slaughter MM. Origin of transient and sustained responses in ganglion cells of the retina. J Neurosci 2000;20:7087–7095. 11. Slaughter MM, Miller RF. 2-Amino-4-phosphonobutyric acid: a new pharmacological tool for retina research. Science 1981;211:182–185. 12. Wu SM, Gao F, Maple BR. Functional architecture of synapses in the inner retina: segregation of visual signals by stratification of bipolar cell axon terminals. J Neurosci 2000;20:4462–4470. 13. Li W, DeVries SH. Bipolar cell pathways for color and luminance vision in a dichromatic mammalian retina. Nat Neurosci 2006;9(5):669–675. 14. Roska B, Werblin F. Vertical interactions across ten parallel, stacked representations in the mammalian retina. Nature 2001;410(6828):583–587. 15. Nelson R, Famiglietti EV Jr, Kolb H. Intracellular staining reveals different levels of stratification for on- and off-center ganglion cells in cat retina. J Neurophys 1978;41(2):472–483. 16. Famiglietti J, Edward V, Kaneko A, Tachibana M. Neuronal architecture of on and off pathways to ganglion cells in carp retina. Science 1977;198:1267–1269. 17. Marchiafava PL, Weiler R. Intracellular analysis and structural correlates of the organization of inputs to ganglion cells in the retina of the turtle. Proc R Soc Lond B 1980;208:103–113. 18. Ichinose T, Shields CR, Lukasiewicz PD. Sodium channels in bipolar cells enhance the light sensitivity of ganglion cells. Invest Ophthalmol Vis Sci 2003;44:E-abstract 1007. 19. Wunk DF, Werblin FS. Synaptic inputs to the ganglion cells in the tiger salamander retina. J Gen Physiol 1979;73:265–286. 20. Dong C, Werblin FS. Temporal contrast enhancement via GABAC feedback at bipolar terminals in the tiger salamander retina. J Neurophys 1998;79:2171–2180. 21. Nirenberg S, Meister M. The light response of retinal ganglion cells is truncated by a displaced amacrine circuit. Neuron 1997;18:637–650. 22. Sagdullaev BT, McCall MA, Lukasiewicz PD. Presynaptic inhibition modulates spillover, creating distinct dynamic response ranges of sensory output. Neuron 2006;50(6):923–935. 23. Higgs MH, Lukasiewicz PD. Glutamate uptake limits synaptic excitation of retinal ganglion cells. J Neurosci 1999;19:3691–3700. 24. Lukasiewicz PD, Lawrence JE, Valentino TL. Desensitizing glutamate receptors shape excitatory synaptic inputs to tiger salamander retinal ganglion cells. J Neurosci 1995;15: 6189–6199.

302

Sagdullaev et al.

25. Lagnado L, Gomis A, Job C. Continuous vesicle cycling in the synaptic terminal of retinal bipolar cells. Neuron 1996;17:957–967. 26. von Gersdorff H, Sakaba T, Berglund K, Tachibana M. Submillisecond kinetics of glutamate release from a sensory synapse. Neuron 1998;21:1177–1188. 27. Sterling P, Matthews G. Structure and function of ribbon synapses. Trends Neurosci 2005;28(1):20–29. 28. Isaacson JS, Nicoll RA. The uptake inhibitor L-trans-PDC enhances responses to glutamate but fails to alter the kinetics of excitatory synaptic currents in the hippocampus. J Neurophysiol 1993;70(5):2187–2191. 29. Sarantis M, Ballerini L, Miller B, Silver RA, Edwards M, Attwell D. Glutamate uptake from the synaptic cleft does not shape the decay of the non-NMDA component of the synaptic current. Neuron 1993;11:541–549. 30. Matsui K, Hosoi N, Tachibana M. Active role of glutamate uptake in the synaptic transmission from retinal nonspiking neurons. J Neurosci 1999;19:6755–6766. 31. Eliasof S, Werblin F. Characterization of the glutmate transporter in retinal cones of the tiger salamander. J Neurosci 1993;13:402–411. 32. Otis TS, Wu YC, Trussell LO. Delayed clearance of transmitter and the role of glutamate transporters at synapses with multiple release sites. J Neurosci 1996;16:1634–1644. 33. Isaacson JS. Glutamate spillover mediates excitatory transmission in the rat olfactory bulb. Neuron 1999;23:377–384. 34. Trussell LO, Thio LL, Zorumski CF, Fischbach GD. Rapid desensitization of glutamate receptors in vertebrate central neurons. Proc Natl Acad Sci U S A 1988;85:2834–2838. 35. Higgs MH, Tran MN, Romano C, Lukasiewicz PD. Group III metabotropic glutamate receptors inhibit glutamate release from retinal bipolar cells. Invest Ophthalmol Vis Sci 2000;41:S621. 36. Awatramani GB, Slaughter MM. Intensity-dependent, rapid activation of presynaptic metabotropic glutamate receptors at a central synapse. J Neurosci 2001;15:741–749. 37. Palmer MJ, Hull C, Vigh J, von Gersdorff H. Synaptic cleft acidification and modulation of short-term depression by exocytosed protons in retinal bipolar cells. J Neurosci 2003;23(36):11332–11341. 38. Lukasiewicz PD, Werblin FS. A novel GABA receptor modulates synaptic transmission from bipolar to ganglion and amacrine cells in the tiger salamander retina. J Neurosci 1994;14:1213–1223. 39. Zhang J, Chang-Sub J, Slaughter MM. Serial inhibitory synapses in retina. Vis Neurosci 1997;14:553–563. 40. Roska B, Nemeth E, Werblin FS. Response to change Is facilitated by a three-neuron disinhibitory pathway in the tiger salamander retina. J Neurosci 1998;18:3451–3459. 41. Eggers ED, Lukasiewicz PD. GABAA, GABAC and glycine receptor-mediated inhibition differentially affects light-evoked signaling from mouse retinal rod bipolar cells. J Physiol 2006;572:215–225. 42. Masland RH. The many roles of starburst amacrine cells. Trends Neurosci 2005;28(8): 395–396. 43. Werblin F, Roska B, Balya D. Parallel processing in the mammalian retina: lateral and vertical interactions across stacked representations. Prog Brain Res 2001;131:229–238. 44. Cook PB, McReynolds JS. Lateral inhibition in the inner retina is important for spatial tuning of ganglion cells. Nat Neurosci 1998;1:714–719. 45. Flores-Herr N, Protti DA, Wassle H. Synaptic currents generating the inhibitory surround of ganglion cells in the mammalian retina. J Neurosci 2001;21:4852–4863. 46. Ichinose T, Lukasiewicz PD. Inner and outer retinal pathways both contribute to surround inhibition of salamander ganglion cells. J Physiol 2005;565(Pt 2):517–535.

Inner Retinal Processing

303

47. Euler T, Wassle H. Different contributions of GABAA and GABAC receptors to rod and cone bipolar cells in a rat retinal slice preparation. J Neurophys 1998;79:1384–1395. 48. Shields CR, Tran MN, Wong RO, Lukasiewicz PD. Distinct ionotropic GABA receptors mediate presynaptic and postsynaptic inhibition in retinal bipolar cells. J Neurosci 2000;20(7):2673–2682. 49. Eggers ED, Lukasiewicz PD. Receptor and transmitter release properties set the time course of retinal inhibition. Journal of Neuroscience 2006;26:9413–9425. 50. Eggers ED, Lukasiewicz PD. Differential inhibitory signaling to bipolar cell classes in mouse retina. Invest Ophthalmol Vis Sci 2005;46(5):1125. 51. Diamond JS, Jahr CE. Asynchronous release of synaptic vesicles determines the time course of the AMPA receptor-mediated EPSC. Neuron 1995;15:1097–1107. 52. Ichinose T, Lukasiewicz PD. GABA transporters regulate inhibition in the retina by limiting GABAC receptor activation. J Neurosci 2002;22:3285–3292. 53. Menger N, Pow DV, Wassle H. Glycinergic amacrine cells of the rat retina. J Comp Neurol 1998;401:34–46. 54. Haverkamp S, Muller U, Harvey K, Harvey RJ, Betz H, Wassle H. Diversity of glycine receptors in the mouse retina: localization of the alpha3 subunit. J Comp Neurol 2003;465(4):524–539. 55. Grunert U, Wassle H. Immunocytochemical localization of glycine receptors in the mammalian retina. J Comp Neurol 1993;335(4):523–537. 56. Ivanova E, Muller U, Wassle H. Characterization of the glycinergic input to bipolar cells of the mouse retina. Eur J Neurosci 2006;23(2):350–364. 57. Nelson R. AII amacrine cells quicken time course of rod signals in the cat retina. J Neurophysiol 1982;47(5):928–947. 58. Dacheux RF, Raviola E. The rod pathway in the rabbit retina: a depolarizing bipolar and amacrine cell. J Neurosci 1986;6(2):331–345. 59. Muller F, Wassle H, Voigt T. Pharmacological modulation of the rod pathway in the cat retina. J Neurophys 1988;59(6):1657–1672. 60. Deans MR, Volgyi B, Goodenough DA, Bloomfield SA, Paul DL. Connexin36 is essential for transmission of rod-mediated visual signals in the mammalian retina. Neuron 2002;36(4):703–712. 61. Rockhill RL, Daly FJ, MacNeil MA, Brown SP, Masland RH. The diversity of ganglion cells in a mammalian retina. J Neurosci 2002;22(9):3831–3843. 62. Enroth-Cugell C, Robson JG. The contrast sensitivity of retinal ganglion cells of the cat. J Physiol (Lond) 1966;187:517–552. 63. Boycott BB, Wassle H. The morphological types of ganglion cells of the domestic cat’s retina. J Physiol (Lond) 1974;240:397–419. 64. Callaway EM. Structure and function of parallel pathways in the primate early visual system. J Physiol 2005;566(Pt 1):13–19. 65. Van Essen DC, Gallant JL. Neural mechanisms of form and motion processing in the primate visual system. Neuron 1994;13(1):1–10. 66. Barlow HB, Hill RM, Levick WR. Retinal ganglion cells responding selectively to direction and speed of image motion in the rabbit. J Physiol (Lond) 1964;173:377–407. 67. Taylor WR, He S, Levick WR, Vaney DI. Dendritic computation of direction selectivity by retinal ganglion cells. Science 2000;289(5488):2347–2350. 68. Fried SI, Munch TA, Werblin FS. Mechanisms and circuitry underlying directional selectivity in the retina. Nature 2002;420(6914):411–414. 69. Euler T, Detwiler PB, Denk W. Directionally selective calcium signals in dendrites of starburst amacrine cells. Nature 2002;418(6900):845–852.

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Sagdullaev et al.

70. Fried SI, Munch TA, Werblin FS. Directional selectivity is formed at multiple levels by laterally offset inhibition in the rabbit retina. Neuron 2005;46(1):117–127. 71. Yoshida K, Watanabe D, Ishikane H, Tachibana M, Pastan I, Nakanishi S. A key role of starburst amacrine cells in originating retinal directional selectivity and optokinetic eye movement. Neuron 2001;30(3):771–780. 72. Berson DM, Dunn FA, Takao M. Phototransduction by retinal ganglion cells that set the circadian clock. Science 2002;295(5557):1070–1073. 73. Masland RH. Neuronal diversity in the retina. Curr Opin Neurobiol 2001;11:431–436. 74. Eggers ED, McCall MA, Lukasiewicz PD. Presynaptic inhibition differentially shapes transmission in distinct circuits in the mouse retina. J Physiol. 2007;582(Pt2):569–82.

14 Human Cone Spectral Sensitivities and Color Vision Deficiencies Andrew Stockman and Lindsay T. Sharpe CONTENTS Introduction Cone Spectral Sensitivities Factors that Influence Spectral Sensitivity Congenital Color Vision Deficiencies Conclusions References

INTRODUCTION Overview A precise knowledge of the spectral sensitivities of the human cones is essential for the understanding and modeling of both normal and defective color vision. Here, we discuss cone spectral sensitivities, their relationship to color matching, and their derivation from psychophysical measurements obtained from normal trichromatic, dichromatic and monochromatic observers. We present a consistent set of mean 2-deg and 10-deg cone fundamentals based on the work of Stockman, Sharpe and Fach [1] and Stockman and Sharpe [2]. Along with the associated lens and macular pigment and photopigment templates, these can be used to model color vision at the cornea and retina and to account for individual differences. The common forms of congenital color vision deficiencies are due to alterations to the spectral sensitivities of the cones, or the loss of one or more cone photopigments. We consider these deficits, their symptoms and diagnosis. Transduction Human spectral sensitivity is determined at the very first step in the phototransduction cascade by the energy required to isomerize the visual chromophore in the rod and cone photoreceptors from 11-cis retinal to all-trans retinal. This isomerization triggers the rapid conformational change of the G protein-coupled-receptor-protein rhodopsin into the activated photoproduct, metarhodopsin II, which then activates transducin, the G protein molecule, by initiating the separation of the α-transducin from the trimer (see [3–8]). From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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The isomerization of the chromophore is energized by the absorption of a photon. The likelihood that a given photon will produce an isomerization depends on how closely its energy matches the energy required for the transition. Although the same chromophore is found in all the human photopigments, the isomerization energy is modified by its environment and, in particular, the identity of key amino acids in the parts of the opsin molecule that surround the chromophore. These modify the isomerization energy and thus the spectral sensitivity of the photoreceptor (for reviews of spectral tuning, see [9–11]). In most observers with normal color vision, there are four photoreceptor classes: three types of cone photoreceptors, which are referred to as long-, middle-, and short-wavelength sensitive (L, M, and S), and a single type of rod photoreceptor. Rods, which are more sensitive than cones, mediate vision at night when photons are relatively scarce, whereas cones mediate color vision during the day when photons are abundant. Those conditions under which the rods and the cones operate alone are known as scotopic and photopic, respectively, while those under which they operate jointly are known as mesopic (see Chapter 15 on luminous efficiency functions). Although the cone spectral sensitivities, at the retina, depend mainly on the energy required to isomerize the chromophore, the spectral sensitivities of the human observer measured behaviorally, at the cornea, also depend on the absorption by the optical media and on the density of photopigment in the photoreceptor outer segment, which vary between observers (see the section on other factors that influence spectral sensitivity). Univariance, Monochromacy, Dichromacy, and Trichromacy When a photon causes the isomerization of the chromophore and triggers the phototransduction cascade, the effect on the output is independent of its wavelength. Thus, photoreceptors are effectively sophisticated photon counters, the outputs of which vary univariantly according to the number of photons that are absorbed (e.g., [12, 13]). If the rate of photon absorption changes, it is impossible to tell from the photoreceptor output whether the changes are due to a variation in light intensity or in wavelength. In other words, color and intensity are confounded, and individual rods or cones are color blind. Thus, human color perception requires comparisons between the activities of more than one type of photoreceptor that have distinct spectral sensitivities. If only one photoreceptor type operates, vision is monochromatic or reduced to a single dimension: Two lights of any spectral composition can be made to match perfectly simply by matching their intensities. All normal observers are monochromatic under scotopic conditions (e.g., dim starlight), when only the rods are functioning. However, some rare human observers are monochromatic under photopic conditions because they have either no operating cone photoreceptors (rod monochromats, discussed in a separate section) or only one of the three types (cone monochromats, also discussed in a separate section). If only two cone photoreceptors operate, vision is dichromatic or reduced to two dimensions: Lights of any spectral composition can be matched by a mixture of two other lights. Human observers, who are dichromatic, fall into three classes: protanopes, deuteranopes, and tritanopes, depending on whether they are missing, respectively, the L, M, or S cones (see the sections on protan and deutan defects and on tritanopia). Most observers with normal color vision have three classes of cone photoreceptor and are therefore trichromatic (i.e., they have three dimensions of color vision).

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Fig. 1. Left: The amounts of each of the 444-, 526-, and 645-nm primaries (the tristimulus values) required to match monochromatic lights spanning the visible spectrum are known as the red (R), green (G), and blue (B) color-matching functions or CMFs (red, green, and blue lines respectively; the wavelengths of the primaries are shown by the vertical dashed lines). A negative sign means that the primary must be added to the target to complete the match. CMFs can be linearly transformed from one set of primaries to another. The only restriction on the choice of primary lights is that they must be independent—in the sense that no two will match the third. Right: CMFs for the imaginary cone fundamental L, M, and S primaries, primary lights that would uniquely stimulate the L, M, and S cones, respectively. The fundamentals are the 10° cone sensitivities of Stockman and Sharpe [2].

Trichromacy and Color-Matching Functions Because normal human photopic vision is trichromatic, the color of any light can be defined or matched by just three variables: the intensities of three specially selected or “independent” primary lights, which are typically chosen to be red (R), green (G), and blue (B) (with the essential proviso that no two will match the third). The left panel of Fig. 1 shows examples of the red, green, and blue color-matching functions (CMFs; also known as tristimulus values) for RGB primaries of 645, 526, and 444 nm. Each CMF defines the amount of that primary required to match monochromatic targets throughout an equal energy visible spectrum. The CMFs can be linearly transformed to any other set of real primary lights and, as illustrated in Fig. 1, to imaginary primary lights, such as the L, M, and S cone fundamental primaries (right panel). These three fundamental primaries (or Grundempfindungen, fundamental sensations) are the three imaginary primary lights that would uniquely stimulate each of the three cones to yield the L-, M-, and S-cone spectral sensitivity functions (such lights are not physically realizable because of the overlapping spectral sensitivities of the cone photopigments). The three CMFs corresponding to the three fundamental primaries are the cone fundamental CMFs or cone spectral sensitivities. For a further discussion of colorimetry and its link to the cone fundamentals, see the work of Stockman and Sharpe [2] and Stockman [14]. A knowledge of the linear transformation from the red, green, and blue CMFs to the three cone fundamental CMFs would allow us to define the mean human cone

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spectral sensitivities, but this information is not directly available from normal color matches (except for the S-cone fundamentals; see [15, 16]). Direct measurements of the cone spectral sensitivities made using special, color-deficient, observers not only yield estimates of the cone spectral sensitivities themselves, but also allow estimates of the transformation from the CMFs (which is sometimes preferable to the raw spectral sensitivity data). CONE SPECTRAL SENSITIVITIES Introduction Since the establishment of trichromatic color theory (e.g., [17–19]), a central goal of vision science has been the accurate determination of the L-, M-, and S- cone spectral sensitivities. Studies of human cone spectral sensitivity have encompassed many fields of inquiry, including fundus reflectometry (e.g., [20]), microspectrophotometry (e.g., [21]), suction electrode recordings (e.g., [22, 23]), electroretinography (e.g., [24]), and absorption spectroscopy [25–28]. Visual psychophysical experiments in human observers still provide the most extensive and accurate spectral sensitivity data. Arguably, the first plausible psychophysical estimates of the cone spectral sensitivities were obtained by König and Dieterici [29] (see symbols, Fig. 3, below). Since then, many other estimates have been made, notably those by Bouma [30]; Judd [31, 32]; Wyszecki and Stiles [33]; Vos and Walraven [34]; Vos [35]; Estévez [36], Vos, Estévez, and Walraven [37]; and Stockman, MacLeod, and Johnson [16]. These have been discussed elsewhere (e.g., [38, 39–41]). Until recently, the estimates by Smith and Pokorny [42] have been widely used in science and research as a de facto standard. However, newer estimates by Stockman and Sharpe [2] have improved on these and have now been proposed as the Commission Internationale de l’ Éclairage (International Lighting Commission, CIE) standard for “physiologically relevant” fundamental primaries (CIE Technical Reports: Fundamental Chromaticity Diagram with Physiological Axes, Parts 1 and 2, CIE, Vienna). Most estimates of the cone spectral sensitivities depend on the use of dichromat observers, missing one of the three normal cone types (see the sections on protanopia and deuteranopia and on tritanopia) and the assumption—known as the “loss”, “reduction,” or “König” hypothesis—that their remaining cone classes are normal [29, 43], an approach that now has a firm foundation, molecular genetically [44, 45] and psychophysically [46]. S-cone (or blue-cone) monochromats [47, 48], which are doubly reduced in the sense that they lack both the L and M cones, are particularly useful for measuring S-cone spectral sensitivity (see the section on cone monochromacies). Cone Spectral Sensitivity Measurements The cone fundamentals can be estimated by comparing dichromatic and normal color matches [19, 49] and then deriving a transformation matrix based on the locations of the confusion points of the three varieties of dichromacy. These confusion or copunctal points are the chromaticities at which the color confusion lines of the dichromats converge (see the section on congenital color vision deficiencies). They correspond to the chromaticities of the missing fundamental primaries.

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Fig. 2. Mean spectral sensitivity data. L-cone data from 17 L(ser180) subjects (filled squares) and 5 L(ala180) subjects (open circles) and M-cone data from 9 L1M2/L2M3 protanopes (gray diamonds) measured by Sharpe et al. [46] and S-cone data from 5 normals and 3 blue cone monochromats (filled triangles) measured by Stockman, Sharpe, and Fach [1].

The most straightforward method of estimating the cone fundamentals, however, is to measure the three cone spectral sensitivities directly. Since the three cone spectral sensitivities overlap extensively throughout the spectrum, trichromatic observers are not particularly useful for such measurements. In them, the isolation of the response of a single cone type over substantial regions of the spectrum requires special procedures to favor the desired cone type and disfavor the two undesired ones. A now classical approach is to use selective chromatic adaptation (e.g., [50, 51]) and to present a target of variable wavelength on a larger adapting or background field of a second wavelength (or mixture of wavelengths) that selectively suppresses the sensitivities of the two unwanted cone types. However, cone isolation in normal observers is still difficult to achieve throughout the spectrum. One way of overcoming this difficulty is to use selective adaptation in genotypically identified dichromats, who lack one of the three cone types. With the S cones disadvantaged or suppressed, L- and M-cone spectral sensitivities can be directly measured in deuteranopes who lack M-cone function and in protanopes without L-cone function. Figure 2 shows the mean spectral sensitivity data obtained from 9 L1M2/ L2M3 protanopes (green diamonds), from 17 single-gene L(ser180) deuteranopes with serine at position 180 of their L-cone photopigment opsin gene (red squares), and from 5 single-gene L(ala180) deuteranopes with alanine at position 180 (open circles) (for further details, see [2, 46]). The two mean L-cone functions, which are separated by about 2.5 nm in λmax [46], reflect the two commonly occurring L-cone photopigment

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Fig. 3. S-, M-, and L-cone 2° spectral sensitivity estimates (short-dashed, long-dashed, and solid lines, respectively) of Stockman and Sharpe [2] based on linear transformations of the Stiles and Birch [58] 10° red, green, blue (RGB) color-matching functions (CMFs) using the mean spectral sensitivity data shown in Fig. 2 as a guide. They are compared with the historical S-, M-, and L-cone estimates of König and Dieterici [29] (filled triangles, gray diamonds, and open squares, respectively). The lower inset shows the mean macular density spectrum for a 2° field (dashed line) based on measurements by Bone, Landrum, and Cains [68] and recommended by Stockman and Sharpe [2, 41] and the mean lens density spectrum of van Norren and Vos [59] and slightly adjusted by Stockman, Sharpe, and Fach [1] (solid line).

polymorphisms (see the section on protan and deutan defects). An overall L-cone mean was also derived (not shown) to reflect the proportions of the two polymorphic variants in the population [2]. Tritanopes, who lack the S cones, are less useful in this context because their closely overlapping L- and M-cone spectral sensitivities are hard to isolate by selective adaptation. S-cone spectral sensitivity is most easily measured throughout the spectrum in S-cone monochromats (e.g., [48, 52–57]). In defining a mean S-cone spectral sensitivity, Stockman, Sharpe, and Fach [1] measured S-cone spectral sensitivities in three blue cone monochromats known to lack L and M cones on genotypical as well as phenotypical grounds and combined them with normals’ S-cone data obtained at short and middle wavelengths on an intense yellow background field that selectively adapted

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the M and L cones. Their mean S-cone function is shown in Fig. 2 (filled triangles). The advantage of using S-cone monochromats is that it allows the S-cone spectral sensitivity to be defined at middle and long wavelengths, where it is occluded in normal observers by the more sensitive responses of the M and L cones even when selective chromatic adaptation is employed. From Cone Spectral Sensitivities to Color-Matching Functions Although the cone spectral sensitivities could be defined as the direct sensitivity measurements shown in Fig. 2, it is customary to define them in terms of linear combinations of a set of CMFs, which are—in principle at least—more precise because they are typically based on more data and because the measurements are less variable. All that is required is to find the linear combinations of red (R), green (G), and blue (B) CMFs that best fit each cone spectral sensitivity, allowing adjustments in the densities of prereceptoral filtering and photopigment optical density to account for differences in the mean densities between different populations (these factors are age and race dependent and highly variable between individuals) and to account for differences in retinal area (because the filtering densities change with retinal eccentricity). For more information, see the next section and Chapter 15 on luminous efficiency functions. Figure 3 shows the current 2° estimates of Stockman and Sharpe [2] (lines) compared with the much earlier estimates obtained more than 120 years ago by König and Dieterici [29] (symbols). The Stockman and Sharpe 2° estimates are based on a transformation of the Stiles and Burch [58] 10° CMFs adjusted to 2°. These 10° CMFs, which were measured in 49 subjects from approximately 390 to 730 nm (and in 9 subjects from 730 to 830 nm), are probably the most secure and accurate set of existing color-matching data.

OTHER FACTORS THAT INFLUENCE SPECTRAL SENSITIVITY Several other factors influence spectral sensitivities and color matches. The most important ones arise from individual differences among observers. They should be taken into account when trying to predict the spectral sensitivities of an individual from standard or mean functions. Some of them vary with retinal position and so should be considered when trying to predict the spectral sensitivities for retinal areas or retinal positions that differ from the centrally viewed 2° or 10° areas used to obtain the standard functions. Lens Pigment Light is brought into focus on the retina by the cornea and the pigmented crystalline lens. The pigment in the lens absorbs light mainly of short wavelengths (see lower inset of Fig. 3, black line). Individual differences in lens pigment density can be large, with a range of approximately ±25% of the mean density in young observers ( 390 nm) of LF-soluble components is 120 times greater than for A2E. Inset: Absorption spectrum of A2E after blowing out the ordinate axis. It needs to be kept in mind that the calculations of the expected absorption of light by lipofuscin extracts and A2E refer to chromophores in solutions. Under physiological conditions, all chromophores are encapsulated within lipofuscin granules; therefore, the absorption cross section at the RPE may be substantially different from that in solution. Wavelength dependence of initial rates of light-induced oxygen uptake normalized to equal number of incident photons (action spectra) for suspension of LF granules (B), insoluble (ILF) and soluble (SLF) components of lipofuscin (C) and A2E in liposomes containing unsaturated lipids (D). Note the rate of photooxidation increases with decreasing wavelength for LF, SLF, and ILF, while for A2E it matches its absorption spectrum. (Modified from [33, 217].)

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resembles the wavelength dependence of lipofuscin photooxidation. Thus, due to photochemical properties of oxidized DHA and its high abundance of DHA in POSs, the precursor material of lipofuscin, it may be speculated that the degradation products of DHA are more likely than the degradation products of A2E to be mainly responsible for mediating photoreactivity of lipofuscin. Importantly, the soluble components of lipofuscin account only for half or less of the total dry mass of lipofuscin granules. The remaining components of lipofuscin also absorb light and exhibit photoreactivity similar to SLF. Moreover, their contribution to overall reactivity of lipofuscin increases with age and is responsible for an age-related increase in photoreactivity of lipofuscin granules [33]. With age, there is progressive accumulation of chromophores in the lens, filtering out all the UV and, later in life, also blue and even partly green light, therefore diminishing the probability of photoexcitation of ATR [136] and, subsequently, photooxidative stress mediated by ATR. Lipofuscin, however, still efficiently absorbs longer-wavelength blue and green light, which can still induce photosensitized generation of singlet oxygen and photooxidation [174, 175] (Fig. 5). In summary, aging increases the risk of lipofuscin-dependent oxidative stress in the outer retina due to age-related accumulation of lipofuscin and an increase in aerobic photoreactivity of a single lipofuscin granule. Toxicity of RPE Lipofuscin Significantly, it has been shown that susceptibility of human RPE cells to lightinduced oxidation increases with age and exhibits similar wavelength dependence of photooxidation as lipofuscin [175]. The phototoxic potential of retinal lipofuscin has been demonstrated in cultured human RPE cells fed lipofuscin granules and exposed to either blue-green light (390–550 nm; 2.8 mW/cm2) or yellow-red light (550–800 nm; 2.8 mW/cm2) [36]. Only cells that were fed lipofuscin granules and exposed to 390- to 550-nm light for 12 h (dose of 121 J/cm2) or more exhibited significant morphological changes, loss of lysosomal integrity, enhanced peroxidation of lipids, and greatly reduced survival. Notably, RPE cells in this experiment were exposed to only 300 lipofuscin granules per cell, and part of the granules was phagocytosed. Lipofuscin accumulation in vivo can greatly exceed that value. As mentioned, based on morphometric study of macular RPE from donors 81 to 90 years old, lipofuscin can occupy as much as 19% of cell volume, which corresponds to almost 8,000 granules per cell [11]. In the following experiments, under conditions of sublethal exposure of cultured RPE cells to lipofuscin and blue-green light, it was demonstrated that lipofuscin inhibits several antioxidant and lysosomal enzymes in the dark, and the effect is exacerbated during exposure to light [176, 177]. The phototoxicity of lipofuscin to RPE cells in culture cannot be explained by the action of A2E. Lipofuscin exerted its toxic effects to the RPE cell in vitro, with the content of A2E in cells at most 13 ng per million cells [36], so it was at least two orders of magnitude lower than that required to elicit detectable damaging effects mediated by A2E in the dark [56] and 17 times lower than that required to elicit A2E-mediated phototoxicity induced by 60-s exposure to blue light of 7.5 W/cm2 irradiance (dose of 450 J/cm2) [178].

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In low concentrations, A2E was shown to actually protect against UV light-induced DNA damage in RPE cells in vitro [179]. In experiments performed by Roberts and colleagues [179], RPE cells were incubated for 2 h in culture medium without or with A2E, after which the cells were washed and exposed to UV light. Cells preincubated in the presence of 5 µM A2E were significantly protected against DNA damage in comparison to the control cells. On the other hand, A2E at high concentration can exert several deleterious effects on cells in the dark and mediate blue-light-induced damage to the RPE cells in vitro (reviewed by [58, 172, 180]). A2E was shown to cause membrane permeabilization, inhibit lysosomal ATPase activity, inhibit cytochrome c oxidase and decrease mitochondrial activity, and induce release of proapoptotic proteins from mitochondria, leading eventually to apoptosis. A2E has been shown to act as a competitive inhibitor of organic anion-transporting protein (OATP) present at the apical microvilli of the rat RPE and to a lesser degree in small retinal vessels. OATP, which is also expressed in the brain and liver, transports amphiphilic compounds across the plasma membrane in a sodiumindependent manner, and in the retina is suspected to be involved in retinoid transport. Most studies on A2E toxicity were done on cells in culture that were exposed to A2E provided as a solution. Under physiological conditions, A2E is believed to be present mainly within the lipofuscin granule, and therefore its toxic action in vitro when it was delivered in solution cannot be extrapolated directly to the situation in vivo. Encapsulation of A2E within the lipofuscin granule, where it accounts for only 0.019–0.024% dry weight, is likely to prevent A2E from exerting its deleterious effects on mitochondria, DNA, and transport proteins. Similar to its precursor ATR, A2E is very susceptible to oxidation. As a result, a variety of epoxides, cyclic peroxides, furanoid oxides, and carbonyl products are formed, and several of these products have been identified in human RPE postmortem, suggesting that they are formed in vivo. It has been shown that products of A2E oxidation delivered in solution to cultured RPE cells can induce oxidative damage to DNA and cytotoxicity. However, being formed within the lipofuscin granule, their toxicity under physiological conditions remains to be shown. As mentioned, ATR plays an essential role in the accumulation of A2E and lipofuscin; however, it was not detected as a lipofuscin component. It may be speculated that retinals and retinols also form epoxides in the retina, but in contrast to A2E, they may more easily diffuse out of the lipofuscin granule and damage cellular components. Interestingly, it seems that retinoids and their derivatives are not essential to propagate photooxidative damage once extensive oxidation of POSs has taken place. Oxidation of isolated POSs by UV light, which induces rapid degradation of all retinoids accompanied by generation of reactive species derived from retinoids and oxygen, and feeding it to cultured RPE cells results in accumulation of autofluorescent material [135, 181, 182]. As a result, the efficiency of phagocytosis decreases, and cells become more susceptible to blue-light-induced toxicity. As mentioned, lipofuscin contains abundant polyunsaturated lipids, oxidation of which results in the formation of toxic 4-hydroxy-nonenal and malondialdehyde, as well as DHA oxidation products of photosensitizing properties. Consistently, our data indicate that oxidized DHA, an abundant component of POSs, exhibits not only toxic properties to cultured RPE cells in the dark, but also its toxicity substantially increases on exposure to blue light [173].

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It is clear that at least in vitro lipofuscin can impose photooxidative stress on the RPE cells, inactivate antioxidant and lysosomal enzymes, enhance lipid peroxidation, and cause lysosomal leakage and loss of cell viability. Although the exact mechanism of lipofuscin-dependent cytotoxicity in the RPE is not known, it is likely that oxidative stress, mediated by the photoexcited lipofuscin, could play an important role. Effects of Lipofuscin Components and Oxidative Stress in the RPE on Proinflammatory and Angiogenic Signaling As discussed, lipofuscin may impose oxidative stress on the RPE. It has been demonstrated on several cell types, including RPE, that oxidative stress can induce proinflammatory and angiogenic signaling [183–185]. For example, a component of lipofuscin, A2E and hydrogen peroxide, a product of irradiation of lipofuscin with light, induces upregulation of a potent proinflammatory factor, cyclooxygenase-2 in ARPE-19 cells, a spontaneously immortalized human RPE cell line [186]. Moreover, cultured ARPE-19 cells under oxidative stress induced by exposure to blue light activate the complement cascade in the human serum overlying the cells, as observed by monitoring the cleavage products of complement component C3, which are known to trigger inflammatory responses [187]. The activation of complement is further increased if cells are exposed to light in the presence of A2E. Moreover, two oxidation products of A2E have been identified, peroxy-A2E and furano-A2E, that lead to a significantly increased level of C3 cleavage products, iC3b. At the same time, the level of iC3b in serum exposed to A2E only remains at the same level as in empty control wells. It is important to determine whether oxidation products of ATR analogous to those of peroxy-A2E and furano-A2E also exhibit the ability of complement activation. These products are expected to be more hydrophilic than peroxy-A2E and furano-A2E and, in contrast to A2E and many A2E oxidation products, not anchored in the lipid membrane. This would facilitate their diffusion out of the POS or lipofuscin granule to reach the choroid. Phagocytosis of oxidized POSs by cultured RPE induces 2.8- and 3.2-fold increased expression of proinflammatory cytokines interleukin 8 (IL-8; also known as the neutrophil chemotactic factor) and MCP-1, respectively, in comparison to cells fed naïve POS [185]. Indeed, IL-8 and MCP-1 are potent chemoattractants for phagocytes and are elevated in the vitreous of patients with retinal neovascularization. Exposure of cultured ARPE-19 cells to oxidative stress induced by tert-butyl hydroperoxide or dl-buthionine-(S,R)-sulfoximine results in a substantial upregulation of gene expression of two isoforms of VEGF, VEGF-A and VEGF-C, and secretion of VEGF-A and VEGF-C proteins [183, 184]. Also, lipofuscin components 4-hydroxy-2-nonenal and A2E/light upregulate VEGF expression [188–191]. VEGF is a potent angiogenic factor that promotes choroidal neovascularization [192]. Also another component of lipofuscin, the CEP–protein adduct, has been found to stimulate angiogenesis in the chick embryo and rat cornea and in a mouse model of laser-induced chorioretinal neovascularization [30, 193]. Interestingly, CEP–protein adducts do not upregulate VEGF in ARPE19 cells in vitro, and administration of antiVEGF antibodies only partly blocks CEP–protein adduct-induced angiogenesis in vivo, while it completely blocks angiogenesis induced by VEGF. These results suggest an involvement of an additional, VEGF-independent, angiogenesis pathway.

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As the RPE is a major source of angiogenic and antiangiogenic factors, dysregulation of the balance between them can contribute to the development of choroidal neovascularization of the retina [192]. In summary, oxidative stress induced by lipofuscin, and certain components of lipofuscin in particular, may contribute to eliciting proinflammatory and angiogenic responses. It is particularly important because AMD is related to chronic low-level inflammation characterized by the presence of inflammatory markers (C-reactive proteins) and—in its so-called wet form—with choroidal neovascularization of the retina [155, 192, 194]. APPROACHES TO DIMINISH LIPOFUSCIN ACCUMULATION OR LIPOFUSCIN-INDUCED DAMAGE Accumulated lipofuscin can be viewed as a proof that RPE cells were exposed to oxidative stress, and once accumulated, lipofuscin can further propagate oxidative damage, in particular when it is excited with short-wavelength visible light. Lipofuscin in the RPE may induce damage to the retina not only directly but also indirectly by inducing proinflammatory and angiogenic signaling, leading to retinal degeneration. Therefore, it is essential to minimize lipofuscin formation and to prevent damage induced by lipofuscin already present. To achieve this, the key element is to minimize ATR accumulation in POS disks and to provide adequate antioxidant protection. ATR accumulation and subsequent lipofuscin accumulation can be successfully diminished by diminishing the exposure to light, by diminishing stores of retinoids in the RPE by depletion of vitamin A, or by pharmacological inhibition of enzymes involved in conversion of all-trans retinyl esters into 11-cis retinal. Depletion of vitamin A, achieved either by dietary depletion or by HPR treatment, needs to be carefully monitored to avoid undesirable effects related to deficiency of its metabolite all-trans retinoic acid—a vital regulator of gene expression and essential factor for maintaining the integrity of the skin and mucous membranes as well as a regulatory factor in the immune system [195]. HRP at concentrations tested in numerous cancer trials induces delayed dark adaptation but otherwise is safe and well tolerated [110]. As mentioned, 13-cis retinoic acid, TDT, and TDH are effective inhibitors of the retinoid cycle and subsequent lipofuscin accumulation. However, as 13-cis retinoic acid acts as a competitive inhibitor, it needs to be present in a high concentration, which can cause teratogenicity and systemic toxicity. The long-term effects of TDT and TDH are unknown. Other inhibitors of isomerization of all-trans retinyl esters are all-trans retinylamine and its isomers and derivatives [196–198]. In addition to direct inhibition of isomerization, all-trans retinylamine is a substrate for LRAT and undergoes reversible acylation to form N-retinylamides [197]. As a result, all-trans retinylamine is a more potent inhibitor of 11-cis retinal synthesis than HPR or 13-cis retinoic acid, when compared at the same doses, and its action is sustained over a period of several days after a single injection. Treatment with all-trans retinylamine effectively prevents light-induced damage to the retina [198]. Therefore, it may be expected that all-trans retinylamine will be effective in prevention of lipofuscin accumulation.

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N-Retinylamine may be metabolized through retinol and retinal into retinoic acid [198]. Importantly, when compared with retinol treatment, only a trace elevation of potentially toxic all-trans retinoic acid is detected. Moreover, N-retinylamine interacts only at micromolar concentrations with retinoic acid receptor and does not activate retinoid X receptor [196]. Yet, treatment with N-retinylamine results in massive changes in gene expression in the liver and retina, similar to those induced by HPR, albeit less dramatic than those induced by 13-cis retinoic acid. Therefore, a thorough study of potential toxicity is required before recommendation of N-retinylamine as a potential prophylactic treatment. Also, there are potential risks related to chronic inhibition of isomerization. The minor drawback of inhibition of 11-cis retinal synthesis is delayed dark adaptation. More important, dysfunction of proteins involved in synthesis of 11-cis retinal, such as RPE65, RDH5, RPE65, LRAT, and CRALBP, results in early-onset photoreceptor degenerations and mild-to-severe loss of vision [103]. It seems that avoiding exposures of the retina to bright short-wavelength light is the simplest preventive measure against ATR accumulation and subsequent oxidative damage mediated by ATR and lipofuscin. Inserting yellow intraocular lenses after cataract surgery can mimic protection offered by the natural lens in the elderly patients [199]. Alternatively, yellow or amber sunglasses may offer the same or even better protection against overexposure to blue light. However, it needs to be remembered that blue light photoactivates melanopsin (maximum at 480 nm) in ganglion cells, which plays an important role in pupillary responses to changing light levels and in setting circadian rhythms [200]. Therefore, chronic filtering out all blue light may have undesirable systemic effects. Certainly, filtering out infrared and short-wavelength light in ophthalmic instruments can decrease the risk of retinal damage during ocular surgery or ophthalmic examination [201, 202], and transient inhibition of the synthesis of 11-cis retinal may offer further protection. As mentioned, the retina is equipped with effective defense mechanisms that, despite harsh conditions putting it at risk of oxidative damage, allow the retina to perform well its function in most cases throughout lifetime. Also, long-term adaptation to the environmental light levels plays an important role in modulating the susceptibility to photooxidative damage and includes decreased concentration of rhodopsin, changes in lipid composition of POSs, and increased concentration of low-molecular antioxidants. To enable this protective mechanism to operate, an adequate dietary intake of antioxidant vitamins and micronutrients is necessary. However, it needs to be pointed out that experiments on animals demonstrated that lifelong exposure to light causes retinal degeneration, with its severity correlating with the intensity of light at which the animals are reared [203]. Moreover, rats reared even at dim cyclic light (∼20–30 lux) exhibit an increasing susceptibility to retinal photodamage with aging, while rats reared in the dark are equally susceptible to light-induced damage at different ages. This suggests that chronic exposure to light results in a gradual decline of efficiency of the defense mechanisms. The retina contains abundant antioxidants that scavenge reactive oxygen species or quench excited states of photosensitizers and singlet oxygen. The retinal antioxidants include low molecular weight antioxidants synthesized endogenously, such as lipoic

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acid and ubiquinone, and of dietary origin (vitamin E, vitamin C, lutein, and zeaxanthin). The retina expresses a number of antioxidant enzymes. Some of the enzymes are directly responsible for decomposition of reactive oxygen species, such as superoxide dismutase, catalase, and glutathione peroxidase, which decompose superoxide, hydrogen peroxide, and lipid hydroperoxides. Other enzymes are responsible for detoxification and removal of toxic products or repair of damaged proteins. For instance, glutathione transferase conjugates secondary products of lipid peroxidation to glutathione, and these adducts can be subsequently excreted from the cell [204]. Several studies in vivo and in vitro have demonstrated that antioxidant protection of the retina can be further improved via antioxidant supplementation or upregulation of antioxidant enzymes [128, 205–209]. For example, loss of photoreceptor RDH activity in the rat retina on exposure to intensive green light may be prevented with a synthetic antioxidant, 1,3-dimethylthiourea [128]. A powerful way to increase the retinal resistance to oxidative stress is exposure to phytochemicals, such as common dietary components (resveratrol, sulforaphane, catechins, flavonoids, allium, and curcumin), which activate cell survival signaling and transcription pathways and inhibit proinflammatory signaling [210, 211]. In particular, sulforaphane, highly abundant in broccoli sprouts, stimulates NRF2-dependent expression of antioxidant and detoxification enzymes such as glutathione, thoredoxin, and NAD(P)H:quinone oxidoreductase and effectively protects RPE cells in vitro and the retina in vivo against photooxidative damage [208, 212, 213]. Last but not least, the retina contains a large concentration of DHA that can, itself or after enzymatic oxidation, effectively inhibit or counteract proapoptotic and inflammatory signaling [214–216]. CONCLUSIONS Phototransduction is necessary for visual perception, but it is inherently related to a release of ATR. There is a great deal of evidence implicating ATR released from opsin as a mediator of acute photodamage to the retina as well as accumulation of lipofuscin in the RPE. Once formed, lipofuscin has the potential to impose further threats to the RPE, which include lipofuscin-mediated oxidative damage, and initiation of signaling pathways, potentially leading to the activation of complement and angiogenesis and therefore contributing to the development of atrophic and neovascular forms of AMD. Therefore, to avoid the risk of oxidative stress and lipofuscin formation, preventive measures against ATR accumulation should be taken, including avoiding overexposure of the retina to bright light and, if necessary, pharmacological intervention inhibiting 11-cis retinal synthesis. As ATR accumulation may be particularly damaging during exposure to light, understanding the signaling pathways responsible for mobilization of all-trans retinyl esters under light adaptation may provide a powerful strategy for prophylactic treatment. With age, the transmission of the short-wavelength light to the retina decreases. However, as both the lipofuscin concentration in the RPE and the photoreactivity of the lipofuscin granule increase with age, the risk of photooxidative damage mediated by lipofuscin remains. To counteract the oxidative stress, minimization of exposure to short-wavelength light and upregulation of antioxidant and detoxifying

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pathways are needed. Recent advances in understanding the molecular pathways leading to elevation of cellular resistance to oxidative damage offer hope that they can be utilized for the benefit of the retina. REFERENCES 1. Terman A, Gustafsson B, Brunk UT. The lysosomal-mitochondrial axis theory of postmitotic aging and cell death. Chem-Biol Interact 2006;163:29–37. 2. Keller JN, Dimayuga E, Chen QH, Thorpe J, Gee J, Ding Q. Autophagy, proteasomes, lipofuscin, and oxidative stress in the aging brain. Int J Biochem Cell Biol 2004;36:2376–2391. 3. Katz ML, Robison WG. What is lipofuscin? Defining characteristics and differentiation from other autofluoreseent lysosomal storage bodies. Arch Gerontol Geriatr 2002;34:169–184. 4. Eldred GE. Lipofuscin and other lysosomal storage deposits in the retinal pigment epithelium. In: Marmor MF, Wolfensberger TJ, eds. The retinal pigment epithelium function and disease. New York: Oxford University Press; 1998:651–668. 5. Yin DZ. Biochemical basis of lipofuscin, ceroid, and age pigment-like fluorophores. Free Radic Biol Med 1996;21:871–888. 6. Stroikin Y, Dalen H, Brunk UT, Terman A. Testing the “garbage” accumulation theory of ageing: mitotic activity protects cells from death induced by inhibition of autophagy. Biogerontology 2005;6:39–47. 7. Sullivan PG, Dragicevic NB, Deng JH, et al. Proteasome inhibition alters neural mitochondrial homeostasis and mitochondria turnover. J Biol Chem 2004;279:20699–20707. 8. Powell SR, Wang P, Divald A, et al. Aggregates of oxidized proteins (lipofuscin) induce apoptosis through proteasome inhibition and dysregulation of proapoptotic proteins. Free Radic Biol Med 2005;38:1093–1101. 9. Sitte N, Huber M, Grune T, et al. Proteasome inhibition by lipofuscin/ceroid during postmitotic aging of fibroblasts. FASEB J 2000;14:1490–1498. 10. Chowdhury PK, Halder M, Choudhury PK, et al. Generation of fluorescent adducts of malondialdehyde and amino acids: Toward an understanding of lipofuscin. Photochem Photobiol 2004;79:21–25. 11. Feeney-Burns L, Hilderbrand ES, Eldridge S. Aging human RPR—morphometric analysis of macular, equatorial, and peripheral cells. Invest Ophthalmol Vis Sci 1984;25:195–200. 12. Delori FC, Goger DG, Dorey CK. Age-related accumulation and spatial distribution of lipofuscin in RPE of normal subjects. Invest Ophthalmol Vis Sci 2001;42:1855–1866. 13. Strauss O. The retinal pigment epithelium in visual function. Physiol Rev 2005;85: 845–881. 14. Weiter JJ, Delori FC, Wing GL, Fitch KA. Retinal pigment epithelial lipofuscin and melanin and choroidal melanin in human eyes. Invest Ophthalmol Vis Sci 1986;27:145–152. 15. von Ruckmann A, Fitzke FW, Bird AC. Distribution of pigment epithelium autofluorescence in retinal disease state recorded in vivo and its change over time. Graefes Arch Clin Exp Ophthalmol 1999;237:1–9. 16. Dorey CK, Wu G, Ebenstein D, Garsd A, Weiter JJ. Cell loss in the aging retina—relationship to lipofuscin accumulation and macular degeneration. Invest Ophthalmol Vis Sci 1989;30:1691–1699. 17. Wing GL, Blanchard GC, Weiter JJ. The topography and age relationship of lipofuscin concentration in the retinal pigment epithelium. Invest Ophthalmol Vis Sci 1978;17:601–607. 18. Birnbach CD, Jarvelainen M, Possin DE, Milam AH. Histopathology and immunocytochemistry of the neurosensory retina in fundus flavimaculatus. Ophthalmology 1994;101: 1211–1219.

452

. . Róz anowski and Róz anowski

19. Delaey JJ, Verougstraete C. Hyperlipofuscinosis and subretinal fibrosis in Stargardts disease. Retin-J Retin Vitr Dis 1995;15:399–406. 20. Lorenz B, Preising MN. Best’s disease. Overview of pathology and its causes. Ophthalmologe 2005;102:111–115. 21. Bergsma DR, Wiggert BN, Funahashi M, Kuwabara T, Chader CJ. Vitamin A receptors in normal and dystrophic human retina. Nature 1977;265:66–67. 22. Kolb H, Gouras P. Electron microscopic observations of human retinitis pigmentosa, dominantly inherited. Invest Ophthalmol 1974;13:487–498. 23. Smith RT, Chan JK, Busuoic M, Sivagnanavel V, Bird AC, Chong NV. Autofluorescence characteristics of early, atrophic, and high-risk fellow eyes in age-related macular degeneration. Invest Ophthalmol Vis Sci 2006;47:5495–5504. 24. Roth F, Bindewald A, Holz FG. Key pathophysiologic pathways in age-related macular disease. Graefes Arch Clin Exp Ophthalmol 2004;242:710–716. 25. Curcio CA. Photoreceptor topography in ageing and age-related maculopathy. Eye 2001; 15:376–383. 26. Kliffen M, VanderSchaft TL, Mooy CM, deJong P. Morphologic changes in age-related maculopathy. Microsc Res Tech 1997;36:106–122. 27. Bazan HEP, Bazan NG, Feeneyburns L, Berman ER. Lipids in human lipofuscin-enriched subcellular-fractions of 2 age populations—comparison with rod outer segments and neural retina. Invest Ophthalmol Vis Sci 1990;31:1433–1443. 28. Schutt F, Bergmann M, Holz FG, Kopitz J. Isolation of intact lysosomes from human RPE cells and effects of A2-E on the integrity of the lysosomal and other cellular membranes. Graefes Arch Clin Exp Ophthalmol 2002;240:983–988. 29. Warburton S, Southwick K, Hardman RM, et al. Examining the proteins of functional retinal lipofuscin using proteomic analysis as a guide for understanding its origin. Mol Vis 2005;11:1122–1134. 30. Gugiu BG, Rozanowska M, Rozanowski B, et al. Proteomic and ultrastructural analyses of human lipofuscin. Invest Ophthalmol Vis Sci 2005;46. 31. Eldred GE, Katz ML. Fluorophores of the human retinal-pigment epithelium—separation and spectral characterization. Exp Eye Res 1988;47:71–86. 32. Eldred GE, Lasky MR. Retinal age pigments generated by self-assembling lysosomotropic detergents. Nature 1993;361:724–726. 33. Rozanowska M, Pawlak A, Rozanowski B, et al. Age-related changes in the photoreactivity of retinal lipofuscin granules: role of chloroform-insoluble components. Invest Ophthalmol Vis Sci 2004;45:1052–1060. 34. Schutt F, Bergmann M, Holz FG, Kopitz J. Proteins modified by malondialdehyde, 4-hydroxynonenal, or advanced glycation end products in lipofuscin of human retinal pigment epithelium. Invest Ophthalmol Vis Sci 2003;44:3663–3668. 35. Sakai N, Decatur J, Nakanishi K, Eldred GE. Ocular age pigment “A2-E”: an unprecedented pyridinium bisretinoid. J Am Chem Soc 1996;118:1559–1560. 36. Davies S, Elliott MH, Floor E, et al. Photocytotoxicity of lipofuscin in human retinal pigment epithelial cells. Free Radic Biol Med 2001;31:256–265. 37. Fishkin NE, Sparrow JR, Allikmets R, Nakanishi K. Isolation and characterization of a retinal pigment epithelial cell fluorophore: an all-trans-retinal dimer conjugate. Proc Natl Acad Sci U S A 2005;102:7091–7096. 38. Fishkin N, Jang YP, Itagaki Y, Sparrow JR, Nakanishi K. A2-Rhodopsin: a new fluorophore isolated from photoreceptor outer segments. Org Biomol Chem 2003;1:1101–1105. 39. Hammer M, Richter S, Guehrs KH, Schweitzer D. Retinal pigment epithelium cell damage by A2-E and its photo-derivatives. Mol Vis 2006;12:1348–1354.

Lipofuscin, Visual Transduction, and Age-Related Changes

453

40. Gaillard ER, Avalle LB, Keller LMM, Wang Z, Reszka KJ, Dillon JP. A mechanistic study of the photooxidation of A2E, a component of human retinal lipofuscin. Exp Eye Res 2004;79:313–319. 41. Avalle LB, Wang Z, Dillon JP, Gaillard ER. Observation of A2E oxidation products in human retinal lipofuscin. Exp Eye Res 2004;78:895–898. 42. Dillon J, Wang Z, Avalle LB, Gaillard ER. The photochemical oxidation of A2E results in the formation of a 5,8,5′,8′-bis-furanoid oxide. Exp Eye Res 2004;79:537–542. 43. Jang YP, Matsuda H, Itagaki Y, Nakanishi K, Sparrow JR. Characterization of peroxy-A2E and furan-A2E photooxidation products and detection in human and mouse retinal pigment epithelial cell lipofuscin. J Biol Chem 2005;280:39732–39739. 44. Wang Z, Keller LMM, Dillon J, Gaillard ER. Oxidation of A2E results in the formation of highly reactive aldehydes and ketones. Photochem Photobiol 2006;82:1251–1257. 45. Docchio F, Boulton M, Cubeddu R, Ramponi R, Barker PD. Age-related-changes in the fluorescence of melanin and lipofuscin granules of the retinal-pigment epithelium—a timeresolved fluorescence spectroscopy study. Photochem Photobiol 1991;54:247–253. 46. Cubeddu R, Docchio F, Ramponi R, Boulton M. Time-resolved fluorescence spectroscopy of the retinal-pigment epithelium—age-related studies. IEEE J Quantum Electron 1990;26:2218–2225. 47. Boulton M, Docchio F, Dayhawbarker P, Ramponi R, Cubeddu R. Age-related-changes in the morphology, absorption and fluorescence of melanosomes and lipofuscin granules of the retinal-pigment epithelium. Vision Res 1990;30:1291–1303. 48. Gaillard ER, Atherton SJ, Eldred G, Dillon J. Photophysical studies on human retinal lipofuscin. Photochem Photobiol 1995;61:448–453. 49. Lamb LE, Zareba M, Plakoudas SN, Sarna T, Simon JD. Retinyl palmitate and the bluelight-induced phototoxicity of human ocular lipofuscin. Arch Biochem Biophys 2001;393: 316–320. 50. Lamb LE, Ye T, Haralampus-Grynaviski NM, et al. Primary photophysical properties of A2E in solution. J Phys Chem B 2001;105:11507–11512. 51. Haralampus-Grynaviski NM, Lamb LE, Clancy CMR, et al. Spectroscopic and morphological studies of human retinal lipofuscin granules. Proc Natl Acad Sci U S A 2003;100: 3179–3184. 52. Delori FC, Dorey CK, Staurenghi G, Arend O, Goger DG, Weiter JJ. In-vivo fluorescence of the ocular fundus exhibits retinal-pigment epithelium lipofuscin characteristics. Invest Ophthalmol Vis Sci 1995;36:718–729. 53. Bindewald-Wittich A, Han M, Schmitz-Valckenberg S, et al. Two-photon-excited fluorescence imaging of human RPE cells with a femtosecond Ti: sapphire laser. Invest Ophthalmol Vis Sci 2006;47:4553–4557. 54. Delori FC. Autofluorescence method to measure macular pigment optical densities fluorometry and autofluorescence imaging. Arch Biochem Biophys 2004;430:156–162. 55. Mata NL, Weng J, Travis GH. Biosynthesis of a major lipofuscin fluorophore in mice and humans with ABCR-mediated retinal and macular degeneration. Proc Natl Acad Sci U S A 2000;97:7154–7159. 56. Sparrow JR, Parish CA, Hashimoto M, Nakanishi K. A2E, a lipofuscin fluorophore, in human retinal pigmented epithelial cells in culture. Invest Ophthalmol Vis Sci 1999;40: 2988–2995. 57. Feeney-Burns L, Berman ER, Rothman H. Lipofuscin of human retinal pigment epithelium. Am J Ophthalmol 1980;90:783–791. 58. Rozanowska M, Sarna T. Light-induced damage to the retina: role of rhodopsin chromophore revisited. Photochem Photobiol 2005;81:1305–1330.

454

. . Róz anowski and Róz anowski

59. Kennedy CJ, Rakoczy PE, Constable IJ. Lipofuscin of the retinal pigment epithelium: a review. Eye 1995;9:763–771. 60. Rodieck RW. The first steps in seeing. Sunderland, MA: Sinauer; 1998. 61. Katz ML, Drea CM, Eldred GE, Hess HH, Robison WG. Influence of early photoreceptor degeneration on lipofuscin in the retinal-pigment epithelium. Exp Eye Res 1986;43:561–573. 62. D’Cruz PM, Yasumura D, Weir J, et al. Mutation of the receptor tyrosine kinase gene Mertk in the retinal dystrophic RCS rat. Hum Mol Genet 2000;9:645–651. 63. Liu JH, Itagaki Y, Ben-Shabat S, Nakanishi K, Sparrow JR. The biosynthesis of A2E, a fluorophore of aging retina, involves the formation of the precursor, A2-PE, in the photoreceptor outer segment membrane. J Biol Chem 2000;275:29354–29360. 64. Katz ML, Eldred GE. Retinal light damage reduces auto-fluorescent pigment deposition in the retinal-pigment epithelium. Invest Ophthalmol Vis Sci 1989;30:37–43. 65. Boulton M, McKechnie NM, Breda J, Bayly M, Marshall J. The formation of auto-fluorescent granules in cultured human RPE. Invest Ophthalmol Vis Sci 1989;30:82–89. 66. Rakoczy P, Kennedy C, Thompsonwallis D, Mann K, Constable I. Changes in retinal-pigment epithelial-cell autofluorescence and protein expression associated with phagocytosis of rod outer segments in vitro. Biol Cell 1992;76:49–54. 67. Wihlmark U, Wrigstad A, Roberg K, Brunk UT, Nilsson SEG. Lipofuscin formation in cultured retinal pigment epithelial cells exposed to photoreceptor outer segment material under different oxygen concentrations. APMIS 1996;104:265–271. 68. Wassell J, Ellis S, Burke J, Boulton M. Fluorescence properties of autofluorescent granules generated by cultured human RPE cells. Invest Ophthalmol Vis Sci 1998;39:1487–1492. 69. Burke JM, Skumatz CMB. Autofluorescent inclusions in long-term postconfluent cultures of retinal pigment epithelium. Invest Ophthalmol Vis Sci 1998;39:1478–1486. 70. Katz ML, Shanker MJ. Development of lipofuscin-like fluorescence in the retinal-pigment epithelium in response to protease inhibitor treatment. Mech Ageing Dev 1989;49:23–40. 71. Katz ML. Incomplete proteolysis may contribute to lipofuscin accumulation in the retinal pigment epithelium. In: Porta EA, ed. Lipofuscin and ceroid pigments. New York: Plenum Press; 1990:109–118. 72. Ivy GO, Kanai S, Ohta M, et al. Lipofuscin-like substances accumulate rapidly in brain, retina and internal organs with cysteine protease inhibition. In: Porta EA, ed. Lipofuscin and ceroid pigments. New York: Plenum Press; 1990:31–47. 73. Rakoczy PE, Zhang D, Robertson T, et al. Progressive age-related changes similar to age-related macular degeneration in a transgenic mouse model. Am J Pathol 2002;161: 1515–1524. 74. Hayasaka S. Aging changes in lipofuscin, lysosomes and melanin in the macular area of human retina and choroid. Jpn J Ophthalmol 1989;33:36–42. 75. Boulton M, Moriarty P, Jarvisevans J, Marcyniuk B. Regional variation and age-relatedchanges of lysosomal-enzymes in the human retinal-pigment epithelium. Br J Ophthalmol 1994;78:125–129. 76. Hoppe G, Marmorstein AD, Pennock EA, Hoff HF. Oxidized low density lipoprotein-induced inhibition of processing of photoreceptor outer segments by RPE. Invest Ophthalmol Vis Sci 2001;42:2714–2720. 77. Hoppe G, O’Neil J, Hoff HF, Sears J. Accumulation of oxidized lipid-protein complexes alters phagosome maturation in retinal pigment epithelium. Cell Mol Life Sci 2004;61: 1664–1674. 78. Crabb JW, O’Neil J, Miyagi M, West K, Hoff HF. Hydroxynonenal inactivates cathepsin B by forming Michael adducts with active site residues. Protein Sci 2002;11:831–840.

Lipofuscin, Visual Transduction, and Age-Related Changes

455

79. Bergmann M, Schutt F, Holz FG, Kopitz J. Inhibition of the ATP-driven proton pump in RPE lysosomes by the major lipofuscin fluorophore A2-E may contribute to the pathogenesis of age-related macular degeneration. FASEB J 2004;18:562–564. 80. Bermann M, Schutt F, Holz FG, Kopitz J. Does A2E, a retinoid component of lipofuscin and inhibitor of lysosomal degradative functions, directly affect the activity of lysosomal hydrolases? Exp Eye Res 2001;72:191–195. 81. Holz FG, Schutt F, Kopitz J, et al. Inhibition of lysosomal degradative functions in RPE cells by a retinoid component of lipofuscin. Invest Ophthalmol Vis Sci 1999;40:737–743. 82. Finnemann SC, Leung LW, Rodriguez-Boulan E. The lipofuscin component A2E selectively inhibits phagolysosomal degradation of photoreceptor phospholipid by the retinal pigment epithelium. Proc Natl Acad Sci U S A 2002;99:3842–3847. 83. Katz ML, Robison WG. Nutritional influences on autooxidation, lipofuscin accumulation, and aging. In: Johnson JEJ, Walford R, Harman D, Miquel J, eds. Free radicals, aging, and degenerative diseases. New York: Liss; 1986:113–129. 84. Boulton M, Rozanowska M, Rozanowski B. Retinal photodamage. J Photochem Photobiol B-Biol 2001;64:144–161. 85. Katz ML, Christianson JS, Gao CL, Handelman GJ. Iron-induced fluorescence in the retina— dependence on vitamin-A. Invest Ophthalmol Vis Sci 1994;35:3613–3624. 86. Hahn P, Qian Y, Dentchev T, et al. Disruption of ceruloplasmin and hephaestin in mice causes retinal iron overload and retinal degeneration with features of age-related macular degeneration. Proc Natl Acad Sci U S A 2004;101:13850–13855. 87. Sundelin SP, Nilsson SEG. Lipofuscin-formation in retinal pigment epithelial cells is reduced by antioxidants. Free Radic Biol Med 2001;31:217–225. 88. Halliwell B, Gutteridge JMC. Free radicals in biology and medicine. 3rd ed. Oxford, UK: Oxford University Press, 2000. 89. Sugano E, Tomita H, Ishiguro SI, Isago H, Tamai M. Nitric oxide-induced accumulation of lipofuscin-like materials is caused by inhibition of cathepsin S. Curr Eye Res 2006;31:607–616. 90. Schadel SA, Heck M, Maretzki D, et al. Ligand channeling within a G-protein-coupled receptor - The entry and exit of retinals in native opsin. J Biol Chem 2003;278:24896–24903. 91. Beharry S, Zhong M, Molday RS. N-Retinylidene-phosphatidylethanolamine is the preferred retinoid substrate for the photoreceptor-specific ABC transporter ABCA4 (ABCR). J Biol Chem 2004;279:53972–53979. 92. Sun H, Nathans J. ABCR, the ATP-binding cassette transporter responsible for Stargardt macular dystrophy, is an efficient target of all-trans-retinal-mediated photooxidative damage in vitro—implications for retinal disease. J Biol Chem 2001;276:11766–11774. 93. Weng J, Mata NL, Azarian SM, Tzekov RT, Birch DG, Travis GH. Insights into the function of Rim protein in photoreceptors and etiology of Stargardt’s disease from the phenotype in abcr knockout mice. Cell 1999;98:13–23. 94. Radu RA, Mata NL, Bagla A, Travis GH. Light exposure stimulates formation of A2E oxiranes in a mouse model of Stargardt’s macular degeneration. Proc Natl Acad Sci U S A 2004;101:5928–5933. 95. Kim SR, Nakanishi K, Itagaki Y, Sparrow JR. Photooxidation of A2-PE, a photoreceptor outer segment fluorophore, and protection by lutein and zeaxanthin. Exp Eye Res 2006;82:828–839. 96. Allikmets R, Singh N, Sun H, et al. A photoreceptor cell-specific ATP-binding transporter gene (ABCR) is mutated in recessive Stargardt macular dystrophy. Nature Genet 1997;15: 236–246. 97. Cremers FPM, van De Pol DJR, van Driel M, et al. Autosomal recessive retinitis pigmentosa and cone-rod dystrophy caused by splice site mutations in the Stargardt’s disease gene ABCR. Hum Mol Genet 1998;7:355–362.

456

. . Róz anowski and Róz anowski

98. Martinez-Mir A, Paloma E, Allikmets R, et al. Retinitis pigmentosa caused by a homozygous mutation in the Stargardt disease gene ABCR. Nat Genet 1998;18:11–12. 99. Zhang QJ, Zulfiqar F, Xiao XS, et al. Severe autosomal recessive retinitis pigmentosa maps to chromosome 1p13.3-p21.2 between D1S2896 and D1S457 but outside ABCA4. Hum Genet 2005;118:356–365. 100. Allikmets R. Further evidence for an association of ABCR alleles with age-related macular degeneration. Am J Hum Genet 2000;67:487–491. 101. Maeda A, Maeda T, Imanishi Y, et al. Role of photoreceptor-specific retinol dehydrogenase in the retinoid cycle in vivo. J Biol Chem 2005;280:18822–18832. 102. Maeda A, Maeda T, Imanishi Y, et al. Retinol dehydrogenase (RDH12) protects photoreceptors from light-induced degeneration in mice. J Biol Chem 2006;281:37697–37704. 103. Travis GH, Golczak M, Moise AR, Palczewski K. Diseases caused by defects in the visual cycle: retinoids as potential therapeutic agents. Annu Rev Pharmocol Toxicol 2007;47: 8.1–8.44. 104. Imanishi Y, Batten ML, Piston DW, Baehr W, Palczewski K. Noninvasive two-photon imaging reveals retinyl ester storage structures in the eye. J Cell Biol 2004;164:373–383. 105. Imanishi Y, Gerke V, Palczewski K. Retinosomes: new insights into intracellular managing of hydrophobic substances in lipid bodies. J Cell Biol 2004;166:447–453. 106. Lamb TD, Pugh EN. Dark adaptation and the retinoid cycle of vision. Prog Retin Eye Res 2004;23:307–380. 107. Bridges CDB, Alvarez RA, Fong SL. Vitamin A in human eyes—amount, distribution, and composition. Invest Ophthalmol Vis Sci 1982;22:706–714. 108. Eldred GE, Katz ML. The auto-fluorescent products of lipid-peroxidation may not be lipofuscin-like. Free Radic Biol Med 1989;7:157–163. 109. Katz ML, Gao CL, Rice LM. Long-term variations in cyclic light intensity and dietary vitamin a intake modulate lipofuscin content of the retinal pigment epithelium. J Neurosci Res 1999;57:106–116. 110. Radu RA, Han Y, Bui TV, et al. Reductions in serum vitamin A arrest accumulation of toxic retinal fluorophores: a potential therapy for treatment of lipofuscin-based retinal diseases. Invest Ophthalmol Vis Sci 2005;46:4393–4401. 111. Jin MH, Li SH, Moghrabi WN, Sun H, Travis GH. Rpe65 is the retinoid isomerase in bovine retinal pigment epithelium. Cell 2005;122:449–459. 112. Moiseyev G, Chen Y, Takahashi Y, Wu BX, Ma JX. RPE65 is the isomerohydrolase in the retinoid visual cycle. Proc Natl Acad Sci U S A 2005;102:12413–12418. 113. Redmond TM, Poliakov E, Yu S, Tsai JY, Lu ZJ, Gentleman S. Mutation of key residues of RPE65 abolishes its enzymatic role as isomerohydrolase in the visual cycle. Proc Natl Acad Sci U S A 2005;102:13658–13663. 114. Wenzel A, Oberhauser V, Pugh EN, et al. The retinal G protein-coupled receptor (RGR) enhances isomerohydrolase activity independent of light. J Biol Chem 2005;280: 29874–29884. 115. Maeda T, Van Hooser JP, Driessen C, Filipek S, Janssen JJM, Palczewski K. Evaluation of the role of the retinal G protein-coupled receptor (RGR) in the vertebrate retina in vivo. J Neurochem 2003;85:944–956. 116. Chen P, Hao WS, Rife L, et al. A photic visual cycle of rhodopsin regeneration is dependent on Rgr. Nature Genet 2001;28:256–260. 117. Winston A, Rando RR. Regulation of isomerohydrolase activity in the visual cycle. Biochemistry 1998;37:2044–2050. 118. McBee JK, Van Hooser JP, Jang GF, Palczewski K. Isomerization of 11-cis-retinoids to alltrans-retinoids in vitro and in vivo. J Biol Chem 2001;276:48483–48493.

Lipofuscin, Visual Transduction, and Age-Related Changes

457

119. Bunt-Milam AH, Saari JC. Immunocytochemical localization of two retinoid-binding proteins in vertebrate retina. J Cell Biol 1983;97:703–712. 120. Saari JC, Bredberg DL. Photochemistry and stereoselectivity of cellular retinaldehydebinding protein from bovine retina. J Biol Chem 1987;262:7618–7622. 121. Kim TS, Maeda A, Maeda T, et al. Delayed dark adaptation in 11-cis-retinol dehydrogenase-deficient mice—a role of RDH11 in visual processes in vivo. J Biol Chem 2005;280: 8694–8704. 122. Saari JC, Bredberg DL, Farrell DF. Retinol esterification in bovine retinal-pigment epithelium—reversibility of lecithin-retinol acyltransferase. Biochem J 1993;291:697–700. 123. Katz ML, Redmond TM. Effect of Rpe65 knockout on accumulation of lipofuscin fluorophores in the retinal pigment epithelium. Invest Ophthalmol Vis Sci 2001;42: 3023–3030. 124. Katz ML, Wendt KD, Sanders DN. RPE65 gene mutation prevents development of autofluorescence in retinal pigment epithelial phagosomes. Mech Ageing Dev 2005;126:513–521. 125. Kim SR, Fishkin N, Kong J, Nakanishi K, Allikmets R, Sparrow JR. Rpe65 Leu450Met variant is associated with reduced levels of the retinal pigment epithelium lipofuscin fluorophores A2E and iso-A2E. Proc Natl Acad Sci U S A 2004;101:11668–11672. 126. Radu RA, Mata NL, Nusinowitz S, Liu XR, Sieving PA, Travis GH. Treatment with isotretinoin inhibits lipofuscin accumulation in a mouse model of recessive Stargardt’s macular degeneration. Proc Natl Acad Sci U S A 2003;100:4742–4747. 127. Maiti P, Kong J, Kim SR, Sparrow JR, Allikmets R, Rando RR. Small molecule RPE65 antagonists limit the visual cycle and prevent lipofuscin formation. Biochemistry 2006;45: 852–860. 128. Darrow RA, Darrow RM, Organisciak DT. Biochemical characterization of cell specific enzymes in light-exposed rat retinas: oxidative loss of all-trans retinol dehydrogenase activity. Curr Eye Res 1997;16:144–151. 129. Schremser JL, Williams TP. Rod outer segment (ROS) renewal as a mechanism for adaptation to a new intensity environment. 1. Rhodopsin levels and ROS length. Exp Eye Res 1995;61:17–23. 130. Reme CE, Wolfrum U, Imsand C, Hafezi F, Williams TP. Photoreceptor autophagy: effects of light history on number and opsin content of degradative vacuoles. Invest Ophthalmol Vis Sci 1999;40:2398–2404. 131. Wiegand RD, Joel CD, Rapp LM, Nielsen JC, Maude MB, Anderson RE. Polyunsaturated fatty acids and vitamin E in rat rod outer segments during light damage. Invest Ophthalmol Vis Sci 1986;27:727–733. 132. Penn JS, Anderson RE. Effects of light history on the rat retina. Prog Retin Res 1991;11: 75–98. 133. Weale RA. Do years or quanta age the retina? Photochem Photobiol 1989;50:429–438. 134. Fite KV, Bengston L, Donaghey B. Experimental light damage increases lipofuscin in the retinal pigment epithelium of Japanese quail (Coturnix coturnix Japonica). Exp Eye Res 1993;57:449–460. 135. Brunk UT, Wihlmark U, Wrigstad A, Roberg K, Nilsson SE. Accumulation of lipofuscin within retinal pigment epithelial cells results in enhanced sensitivity to photooxidation. Gerontology 1995;41:201–211. 136. Boettner EA, Wolter JR. Transmission of the ocular media. Invest Ophthalmol 1962;1: 776–783. 137. Karan G, Lillo C, Yang Z, et al. Lipofuscin accumulation, abnormal electrophysiology, and photoreceptor degeneration in mutant ELOVL4 transgenic mice: a model for macular degeneration. Proc Natl Acad Sci U S A 2005;102:4164–4169.

458

. . Róz anowski and Róz anowski

138. Marmorstein AD, Stanton JB, Yocom J, et al. A model of best vitelliform macular dystrophy in rats. Invest Ophthalmol Vis Sci 2004;45:3733–3739. 139. Nandrot EE, Kim YH, Brodie SE, Huang XZ, Sheppard D, Finnemann SC. Loss of synchronized retinal phagocytosis and age-related blindness in mice lacking alpha v beta 5 integrin. J Exp Med 2004;200:1539–1545. 140. Miceli MV, Newsome DA, Tate DJ, Sarphie TG. Pathologic changes in the retinal pigment epithelium and Bruch’s membrane of fat-fed atherogenic mice. Curr Eye Res 2000;20:8–16. 141. Tanaka N, Ikawa M, Mata NL, Verma IM. Choroidal neovascularization in transgenic mice expressing prokineticin 1: an animal model for age-related macular degeneration. Mol Ther 2006;13:609–616. 142. Kuziel WA, Morgan SJ, Dawson TC, et al. Severe reduction in leukocyte adhesion and monocyte extravasation in mice deficient in CC chemokine receptor 2. Proc Natl Acad Sci U S A 1997;94:12053–12058. 143. Lu B, Rutledge BJ, Gu L, et al. Abnormalities in monocyte recruitment and cytokine expression in monocyte chemoattractant protein 1-deficient mice. J Exp Med 1998;187: 601–608. 144. Ambati J, Anand A, Fernandez S, et al. An animal model of age-related macular degeneration in senescent Ccl-2-or Ccr-2-deficient mice. Nat Med 2003;9:1390–1397. 145. Majji AB, Cao JT, Chang KY, et al. Age-related retinal pigment epithelium and Bruch’s membrane degeneration in senescence-accelerated mouse. Invest Ophthalmol Vis Sci 2000;41:3936–3942. 146. Yu K, Cui YY, Hartzell HC. The bestrophin mutation A243V, linked to adult-onset vitelliform macular dystrophy, impairs its chloride channel function. Invest Ophthalmol Vis Sci 2006;47:4956–4961. 147. Schatz P, Klar J, Andreasson S, Ponjavic V, Dahl N. Variant phenotype of best vitelliform macular dystrophy associated with compound heterozygous mutations in VMD2. Ophthalmic Genet 2006;27:51–56. 148. Marmorstein LY, Wu J, McLaughlin P, et al. The light peak of the electroretinogram is dependent on voltage-gated calcium channels and antagonized by bestrophin (Best-1). J Gen Physiol 2006;127:577–589. 149. Strauss O, Rosenthal R. Function of bestrophin. Ophthalmologe 2005;102:122–126. 150. Rosenthal R, Bakall B, Kinnick T, et al. Expression of bestrophin-1, the product of the VMD2 gene, modulates voltage-dependent Ca2+ channels in retinal pigment epithelial cells. FASEB J 2005;19:178–180. 151. Hartzell C, Putzier I, Arreola J. Calcium-activated chloride channels. Annu Rev Physiol 2005;67:719–758. 152. Ghosh S, Kewalramani G, Yuen G, et al. Induction of mitochondrial nitrative damage and cardiac dysfunction by chronic provision of dietary omega-6 polyunsaturated fatty acids. Free Radic Biol Med 2006;41:1413–1424. 153. Bazan NG. Cell survival matters: docosahexaenoic acid signaling, neuroprotection and photoreceptors. Trends Neurosci 2006;29:263–271. 154. Massaro M, Habib A, Lubrano L, et al. The omega-3 fatty acid docosahexaenoate attenuates endothelial cyclooxygenase-2 induction through both NADP(H) oxidase and PKC epsilon inhibition. Proc Natl Acad Sci U S A 2006;103:15184–15189. 155. Gehrs KM, Anderson DH, Johnson LV, Hageman GS. Age-related macular degeneration - emerging pathogenetic and therapeutic concepts. Ann Med 2006;38:450–471. 156. Valko M, Leibfritz D, Moncol J, Cronin MTD, Mazur M, Telser J. Free radicals and antioxidants in normal physiological functions and human disease. Int J Biochem Cell Biol 2007;39:44–84.

Lipofuscin, Visual Transduction, and Age-Related Changes

459

157. Bui TV, Han Y, Radu RA, Travis GH, Mata NL. Characterization of native retinal fluorophores involved in biosynthesis of A2E and lipofuscin-associated retinopathies. J Biol Chem 2006;281:18112–18119. 158. Mata NL, Tzekov RT, Liu XR, Weng J, Birch DG, Travis GH. Delayed. dark-adaptation and lipofuscin accumulation in abcr+/− mice: implications for involvement of ABCR in age-related macular degeneration. Invest Ophthalmol Vis Sci 2001;42:1685–1690. 159. Polyakov NE, Leshina TV, Konovalova TA, Kispert LD. Carotenoids as scavengers of free radicals in a Fenton reaction: antioxidants or pro-oxidants? Free Radic Biol Med 2001;31:398–404. 160. Rozanowska M, Boulton M, Edge R, et al. Protective effects of phosphatidylethanolamine against retinal reactivity. Free Radic Biol Med 2006;41:S168. 161. Ishii T, Kumazawa S, Sakurai T, Nakayama T, Uchida K. Mass spectroscopic characterization of protein modification by malondialdehyde. Chem Res Toxicol 2006;19:122–129. 162. Uchida K. 4-Hydroxy-2-nonenal: a product and mediator of oxidative stress. Prog Lipid Res 2003;42:318–343. 163. Gu XR, Sun MJ, Gugiu B, Hazen S, Crabb JW, Salomon RG. Oxidatively truncated docosahexaenoate phospholipids: total synthesis, generation, and peptide adduction chemistry. J Org Chem 2003;68:3749–3761. 164. Bernoud-Hubac N, Roberts LJ. Identification of oxidized derivatives of neuroketals. Biochemistry 2002;41:11466–11471. 165. Bernoud-Hubac N, Davies SS, Boutaud O, Montine TJ, Roberts LJ. Formation of highly reactive gamma-ketoaldehydes (neuroketals) as products of the neuroprostane pathway. J Biol Chem 2001;276:30964–30970. 166. Tate DJ, Miceli MV, Newsome DA. Phagocytosis and H2O2 induce catalase and metallothione in gene-expression in human retinal-pigment epithelial-cells. Invest Ophthalmol Vis Sci 1995;36:1271–1279. 167. Miceli MV, Liles MR, Newsome DA. Evaluation of oxidative processes in human pigment epithelial-cells associated with retinal outer segment phagocytosis. Exp Cell Res 1994;214:242–249. 168. Kindzelskii AL, Elner VM, Elner SG, Yang DL, Hughes BA, Petty HR. Human, but not bovine, photoreceptor outer segments prime human retinal pigment epithelial cells for metabolic activation and massive oxidant release in response to lipopolysaccharide and interferon-gamma. Exp Eye Res 2004;79:431–435. 169. Katz ML, Rice LM, Gao CL. Reversible accumulation of lipofuscin-like inclusions in the retinal pigment epithelium. Invest Ophthalmol Vis Sci 1999;40:175–181. 170. Schmitz-Valckenberg S, Bindewald-Wittich A, Dolar-Szczasny J, et al. Correlation between the area of increased autofluorescence surrounding geographic atrophy and disease progression in patients with AMD. Invest Ophthalmol Vis Sci 2006;47:2648–2654. 171. Hwang JC, Chan JWK, Chang S, Smith RT. Predictive value of fundus autofluorescence for development of geographic atrophy in age-related macular degeneration. Invest Ophthalmol Vis Sci 2006;47:2655–2661. 172. Lamb LE, Simon JD. A2E: A component of ocular lipofuscin. Photochem Photobiol 2004;79:127–136. 173. Bakker L, Pawlak A, Rozanowski B, Boulton M, Rozanowska M. Phototoxicity of peroxidized docosahexaenoate. Free Radic Biol Med 2006;41:S156. 174. Rozanowska M, Wessels J, Boulton M, et al. Blue light-induced singlet oxygen generation by retinal lipofuscin in non-polar media. Free Radic Biol Med 1998;24:1107–1112. 175. Rozanowska M, Jarvis-Evans J, Korytowski W, Boulton ME, Burke JM, Sarna T. Blue light-induced reactivity of retinal age pigment—in-vitro generation of oxygen-reactive species. J Biol Chem 1995;270:18825–18830.

460

. . Róz anowski and Róz anowski

176. Wassell J, Davies S, Bardsley W, Boulton M. The photoreactivity of the retinal age pigment lipofuscin. J Biol Chem 1999;274:23828–23832. 177. Shamsi FA, Boulton M. Inhibition of RPE lysosomal and antioxidant activity by the age pigment lipofuscin. Invest Ophthalmol Vis Sci 2001;42:3041–3046. 178. Sparrow JR, Nakanishi K, Parish CA. The lipofuscin fluorophore A2E mediates blue light-induced damage to retinal pigmented epithelial cells. Invest Ophthalmol Vis Sci 2000;41:1981–1989. 179. Roberts JE, Kukielczak BM, Hu DN, et al. The role of A2E in prevention or enhancement of light damage in human retinal pigment epithelial cells. Photochem Photobiol 2002;75:184–190. 180. Sparrow JR, Boulton M. RPE lipofuscin and its role in retinal-pathobiology. Exp Eye Res 2005;80:595–606. 181. Wihlmark U, Wrigstad A, Roberg K, Nilsson SEG, Brunk UT. Lipofuscin accumulation in cultured retinal pigment epithelial cells causes enhanced sensitivity to blue light irradiation. Free Radic Biol Med 1997;22:1229–1234. 182. Sundelin S, Wihlmark U, Nilsson SEG, Brunk UT. Lipofuscin accumulation in cultured retinal pigment epithelial cells reduces their phagocytic capacity. Curr Eye Res 1998;17:851–857. 183. Kannan R, Zhang N, Sreekumar PG, et al. Stimulation of apical and basolateral vascular endothelial growth factor-A and vascular endothelial growth factor-C secretion by oxidative stress in polarized retinal pigment epithelial cells. Mol Vis 2006;12:1649–1659. 184. Sreekumar PG, Kannan R, de Silva AT, Burton R, Ryan SJ, Hinton DR. Thiol regulation of vascular endothelial growth factor-A and its receptors in human retinal pigment epithelial cells. Biochem Biophys Res Commun 2006;346:1200–1206. 185. Higgins GT, Wang JH, Dockery P, Cleary PE, Redmond HP. Induction of angiogenic cytokine expression in cultured RPE by ingestion of oxidized photoreceptor outer segments. Invest Ophthalmol Vis Sci 2003;44:1775–1782. 186. Lukiw WJ, Mukherjee PK, Cui JG, Bazan NG. A2E selectively induces COX-2 in ARPE19 and human neural cells. Curr Eye Res 2006;31:259–263. 187. Zhou JL, Jang YP, Kim SR, Sparrow JR. Complement activation by photooxidation products of A2E, a lipofuscin constituent of the retinal pigment epithelium. Proc Natl Acad Sci U S A 2006;103:16182–16187. 188. Ayalasomayajula SP, Kompella UB. Induction of vascular endothelial growth factor by 4hydroxynonenal and its prevention by glutathione precursors in retinal pigment epithelial cells. Eur J Pharmacol 2002;449:213–220. 189. Yanagi Y, Inoue Y, Iriyama A, Jang WD. Effects of yellow intraocular lenses on lightinduced upregulation of vascular endothelial growth factor. J Cataract Refract Surg 2006;32: 1540–1544. 190. Zhou JL, Cai BL, Jang YP, Pachydaki S, Schmidt AM, Sparrow JR. Mechanisms for the induction of HNE- MDA- and AGE-adducts, RAGE and VEGF in retinal pigment epithelial cells. Exp Eye Res 2005;80:567–580. 191. Schlingemann RO. Role of growth factors and the wound healing response in age-related macular degeneration. Graefes Arch Clin Exp Ophthalmol 2004;242:91–101. 192. Witmer AN, Vrensen G, Van Noorden CJF, Schlingemann RO. Vascular endothelial growth factors and angiogenesis in eye disease. Prog Retin Eye Res 2003;22:1–29. 193. Ebrahem Q, Renganathan K, Sears J, et al. Carboxyethylpyrrole oxidative protein modifications stimulate neovascularization: Implications for age-related macular degeneration. Proc Natl Acad Sci U S A 2006;103:13480–13484. 194. Donoso LA, Kim D, Frost A, Callahan A, Hageman G. The role of inflammation in the pathogenesis of age-related macular degeneration. Surv Ophthalmol 2006;51:137–152.

Lipofuscin, Visual Transduction, and Age-Related Changes

461

195. Blomhoff R, Blomhoff HK. Overview of retinoid metabolism and function. J Neurobiol 2006;66:606–630. 196. Golczak M, Kuksa V, Maeda T, Moise AR, Palczewski K. Positively charged retinoids are potent and selective inhibitors of the trans-cis isomerization in the retinoid (visual) cycle. Proc Natl Acad Sci U S A 2005;102:8162–8167. 197. Golczak M, Imanishi Y, Kuksa V, Maeda T, Kubota R, Palczewski K. Lecithin: retinol acyltransferase is responsible for amidation of retinylamine, a potent inhibitor of the retinoid cycle. J Biol Chem 2005;280:42263–42273. 198. Maeda A, Maeda T, Golczak M, et al. Effects of potent inhibitors of the retinoid cycle on visual function and photoreceptor protection from light damage in mice. Mol Pharmacol 2006;70:1220–1229. 199. Mainster MA. Violet and blue light blocking intraocular lenses: photoprotection versus photoreception. Br J Ophthalmol 2006;90:784–792. 200. Peirson S, Foster RG. Melanopsin: another way of signaling light. Neuron 2006;49: 331–339. 201. Michels M, Sternberg P. Operating microscope-induced retinal phototoxicity—pathophysiology, clinical manifestations and prevention. Surv Ophthalmol 1990;34:237–252. 202. Mainster MA, Ham WT, Delori FC. Potential retinal hazards—instrument and environmental light sources. Ophthalmology 1983;90:927–932. 203. Organisciak DT, Darrow RM, Barsalou L, et al. Light history and age-related changes in retinal light damage. Invest Ophthalmol Vis Sci 1998;39:1107–1116. 204. Maeda A, Crabb JW, Palczewski K. Microsomal glutathione S-transferase 1 in the retinal pigment epithelium: protection against oxidative stress and a potential role in aging. Biochemistry 2005;44:480–489. 205. Gosbell AD, Stefanovic N, Scurr LL, et al. Retinal light damage: Structural and functional effects of the antioxidant glutathione peroxidase-1. Invest Ophthalmol Vis Sci 2006;47:2613–2622. 206. Hanneken A, Lin FF, Johnson J, Maher P. Flavonoids protect human retinal pigment epithelial cells from oxidative-stress-induced death. Invest Ophthalmol Vis Sci 2006;47:3164–3177. 207. Voloboueva LA, Liu JK, Suh JH, Ames BN, Miller SS. (R)-alpha-Lipoic acid protects retinal pigment epithelial cells from oxidative damage. Invest Ophthalmol Vis Sci 2005;46: 4302–4310. 208. Gao XQ, Talalay P. Induction of phase 2 genes by sulforaphane protects retinal pigment epithelial cells against photooxidative damage. Proc Natl Acad Sci U S A 2004;101: 10446–10451. 209. Wrona M, Rozanowska M, Sarna T. Zeaxanthin in combination with ascorbic acid or alphatocopherol protects ARPE-19 cells against photosensitized peroxidation of lipids. Free Radic Biol Med 2004;36:1094–1101. 210. Mattson MP, Cheng AW. Neurohormetic phytochemicals: low-dose toxins that induce adaptive neuronal stress responses. Trends Neurosci 2006;29:632–639. 211. Surh YJ, Kundu JK, Na HK, Lee JS. Redox-sensitive transcription factors as prime targets for chemoprevention with anti-inflammatory and antioxidative phytochemicals. J Nutr 2005;135:2993S–3001S. 212. Tanito M, Masutani H, Kim YC, Nishikawa M, Ohira A, Yodoi J. Sulforaphane induces thioredoxin through the antioxidant-responsive element and attenuates retinal light damage in mice. Invest Ophthalmol Vis Sci 2005;46:979–987. 213. Zhou JL, Gao XQ, Cai BL, Sparrow JR. Indirect antioxidant protection against photooxidative processes initiated in retinal pigment epithelial cells by a lipofuscin pigment. Rejuv Res 2006;9:256–263.

462

. . Róz anowski and Róz anowski

214. Bazan NG. Survival signaling in retinal pigment epithelial cells in response to oxidative stress: significance in retinal degenerations. Adv Exp Med Biol 2006;531–540. 215. Bazan NG. Neuroprotectin D1 (NPD1): a DHA-derived mediator that protects brain and retina against cell injury-induced oxidative stress. Brain Pathol 2005;15:159–166. 216. SanGiovanni JP, Chew EY. The role of omega-3 long-chain polyunsaturated fatty acids in health and disease of the retina. Prog Retin Eye Res 2005;24:87–138. 217. Pawlak A, Rozanowska M, Zareba M, Lamb LE, Simon JD, Sarna T. Action spectra for the photoconsumption of oxygen by human ocular lipofuscin and lipofuscin extracts. Arch Biochem Biophys 2002;403:59–62.

21 A Nonspecific System Provides Nonphotic Information for the Biological Clock Marian H. Lewandowski CONTENTS Introduction Nonphotic Information Nonspecific Systems Intergeniculate Leaflet of the Thalamus Conclusions References

INTRODUCTION Rhythmicity is a key property of all living matter, starting with unicellular organisms and ending with such complex organisms as humans. All the time, numberless structures and functions of the organism undergo rhythmic changes (of seconds, minutes, and hours; circadian; seasonal). These alterations are of great adaptive significance to the organism. They permit behavioral patterns and physiological processes of the organism to be optimally synchronized with the cyclically changing environmental conditions. In recent years, a growing interest in biological rhythms, especially in their mechanism, has been an effect of not only their endogenous origin and the discovery of genetic basis, but—above all—also of the understanding of their great significance for the proper functioning of the organism. One of the first symptoms of functional disturbances (illnesses) in a living organism is a change in the rhythmicity of its physiological processes and behavioral patterns, while interference with the rhythmic processes may be very dangerous and cause disturbances in the functioning of human organism. Therefore, the understanding of the mechanism regulating rhythmic processes, the elements of this mechanism, their reciprocal functional relations, as well as the influence of external factors on its functioning not only presents a challenge to chronobiologists, but also requires specialist knowledge on the part of physicians, pharmacologists, and psychologists, that is, all those in everyday contact with ill and healthy persons. Such information is also valuable to every one of us who listens intently to one’s own organism and wants to live in harmony with it. From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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The discovery of the suprachiasmatic nuclei (SCNs) of the anterior hypothalamus, the main neuronal element building up the mechanism of mammalian biological clock, at the beginning of the 1970s [1] was undoubtedly a giant step in chronobiological research. A direct connection between ganglion cells of the retina and the SCN via the retinohypothalamic tract (RHT) supported the notion of a significant role of light as an external synchronizer (zeitgeber, time donor) of mammalian biological clock [2, 3]. NONPHOTIC INFORMATION However, light is not the only external stimulus that synchronizes the functioning of mammalian biological clock. Also other, nonphotic information has substantial influence on its work [4]. It is of particular significance to higher organisms, including first of all humans, for whom the cyclically changing light/darkness or ambient temperature are not the only factors affecting the normal course of rhythmic processes. Of equal importance as a stimulus synchronizing rhythmic processes is all nonphotic information, which under certain conditions may exert even a more powerful effect than light [5]. In social animals, including mammals above all, there is a strong tendency to synchronize the behavior of an individual with that of other group members. This is particularly important and clearly visible in primates, including humans first, for whom the social instructions developed by way of evolution have become dominant over physicochemical stimuli in the regulation of their behavior [6]. The contemporary lifestyle strongly emphasizes this tendency by showing how far light, present in our everyday activities (shift work, intercontinental flights, etc.), diverges from the solar cycle of day and night, often supplying the biological clock with contradictory and incorrect information about time. A vast body of evidence indicates that synchronization of the biological clock may also take place in the absence of light. Persons kept under permanent poor lighting in specially isolated living quarters or during the polar night show a free-running rhythm, characteristic of the so-called permanent light conditions. In such circumstances, cognitive stimuli themselves and the information about the passing time are not sufficient to change this rhythm. However, the imposition of definite behavioral patterns or the maintenance of regular contacts with the remaining group members may synchronize a free-running rhythm to the circadian rhythm. A similar mechanism can be observed in blind persons, whose acquired permanent behavioral patterns inhibit generation of a free-running rhythm by synchronizing their biological clock to the circadian rhythm. The disturbance of customary everyday behaviors or their elimination as well as deprivation of other social signals may have a serious negative effect on our well-being. A change in mood is often an effect of sudden, unexpected, mainly negative, nonphotic stimuli on the circadian mechanism of the biological clock. Due to their force and negative character, these stimuli can mask the current synchronizing effect of less-potent nonphotic information (synchronizers), thereby causing desynchronization of rhythmic processes. The disturbance of circadian rhythms that frequently accompanies depression may also be connected with behavioral dysfunction, including locomotor ones or changes in customary behaviors, which hinder the availability of social stimuli and thus their synchronizing influence. Nonphotic communication is of particular importance

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between mother and fetus or a newborn. It prepares and optimizes its later behaviors and activities for circadian light conditions [7]. These examples permit the following conclusion: Of importance equal to photic information in man is the direct influence of strictly determined changes and behavioral activity patterns generated by nonphotic stimuli. This is a key factor synchronizing biological rhythms. NONSPECIFIC SYSTEMS A characteristic feature of the central nervous system is its distinct division into two autonomously separate but functionally strictly related systems. The first consists of specific pathways along which there flow centripetal sensory information and centrifugal motor input. These systems are characterized by the selectivity of the flowing information, which indicates that only information specific to a given projection (e.g., optic to the visual cortex or photic to elements of the mechanism of biological clock, including the SCN above all) flows along a definite path. The other system contains nonspecific pathways along which no specific sensory information flows from a particular reception zone or a sense organ to a specific area of the cerebral cortex. Anatomically, they form greatly “disseminated,” diffusible projections with terminals sited in many brain areas, also including the neuronal elements of the biological clock. The activity of sources of these systems causes a general excitation of the brain, the so-called arousal reaction. This arousal, with an electrophysiological picture that is represented by desynchronization of the activity of cortical neurons, makes the information flowing along specific pathways to be well “understood” by the appropriate areas of the brain, mainly the cerebral cortex. Hence, the nonspecific systems prepare the brain for the reception and adequate reaction to stimuli of diverse modality, coming in from specific systems. Only proper “cooperation” of the two systems guarantees normal brain activity—in the context not only of the cortical facilitation of sensory information transmission along nervous pathways but also of their participation in the mechanism of numerous important physiological processes, including the mechanism of mammalian biological clock. Nonphotic stimuli such as behavioral arousal that induces hyperactivity and sleep deprivation, food shortage, as well as such social interactions as, for example, matingoriented behaviors, have no specific projections of their own in the brain. A physiological result of their influence on human organism may be and is activation of nonspecific systems and a change in human brain arousal [7], which also elevate the physical activity of man [4]. They also modulate the work of neuronal elements of the mechanism of the mammalian biological clock, in particular those receiving direct photic information from ganglion cells of the retina. Ascending Reticular-Activating System In 1949, Moruzzi and Magoun [8] demonstrated the indispensability of the reticular formation of the brain stem to the maintenance of cortical activity and behavioral arousal, giving that projection the name ascending reticular-activating system (ARAS). The ascending nonspecific projections of the reticular formation reach brain cortical areas by the dorsal pathway via the thalamus and by the ventral pathway via the basal

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Fig. 1. The sagittal section through the rat brain illustrating the ascending reticular-activating system (ARAS). BF basal forebrain, IGL intergeniculate leaflet, LC locus coeruleus, LDT laterodorsal tegmental nuleus, PPT penduculopontine tegmental nucleus, RN raphe nuclei, SCN suprachiasmatic nuclei, TH thalamus, TMN tuberomammillary nucleus, VTA/SN ventral tegmental area/substantia nigra.

forebrain and the hypothalamus, while neurons of the descending projection of the reticular formation come up to peripheral muscles through activation of motor neurons of the medulla (Fig. 1). The main neurotransmitter of the diffusible thalamocortical projection and some neurons of the basal forebrain is glutamate (GLU); its activity is modified by such neuromodulators of the brain stem as noradrenaline of the locus coeruleus, dopamine of the substantia nigra and ventral tegmental area, serotonin of the raphe nuclei, acetylcholine of pontomesencephalic neurons, or histamine of tuberomammillary neurons. The majority of these projections also have their terminals in the neuronal elements of the mammalian biological clock, receiving information directly or indirectly via basal nuclei of the forebrain [9]. The dominating and most significant projection for the clock activity is the serotoninergic projection from the raphe nuclei, often defined as a nonphotic projection [10]. Orexin/Hypocretin Projection At the end of the 1990s, a group of researchers from San Diego, headed by Gregory Sutcliffe, discovered in the hypothalamic area a peptide controlling appetite and regulating body weight. In earlier studies, rats with lesioned median hypothalamus showed obesity, while those with lesion of the lateral hypothalamus displayed anorexia [11]. The above-mentioned peptide was named hypocretin (Hctr) after the site of its hypothalamic localization and the structural resemblance to the gut hormone secretin [12]. At the same time, in their search for orphan receptor ligands, Yanagisawa’s research group from Texas found two peptides binding to those receptors, orexin A and orexin B. The name orexin (OX) is derived from the Greek word for appetite; it was introduced after an increased appetite had been observed following administration of that peptide into

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the lateral ventricle of rat brain. It shortly turned out that hypocretin and orexin were the same compound participating in the regulation of not only food intake, but also behaviors and behavioral states of the organism, including regulation of the sleep-andwakefulness rhythm. Its concentration changes according to the day-and-night rhythm, reaching its maximum in the waking state and during rapid-eye movement (REM) sleep and the minimum during slow-wave sleep [13]. The blockade of orexin synthesis via degeneration of orexin neurons or genetic mutations causes narcolepsy in animals and humans [14]. To date, two orexin/hypocretin peptides, OX-A (Hctr-1) and OX-B (Hctr-2), have been recognized. They bind to two different metabotropic receptors, OX-R1 and OX-R2. OX-A has high affinity for both these receptors, while OX-B shows considerably greater affinity for the latter receptor (type 2) [15]. The peptide under description is synthesized mainly by a small group of cells of the posterior lateral hypothalamus [16]. A tiny number of small Hctr neurons were also observed in other brain regions: the lateral part of the amygdala, the anterolateral area of the bed nucleus and lateral ventricle [17], as well as in olfactory neurons [18]. Axons of hypocretin neurons, mainly those localized within the hypothalamic area, innervate numerous regions of the brain. The richest projections reach the locus coeruleus and raphe nuclei, which have descending fibers that reach motor neurons, controlling muscle tone, while the ascending fibers that innervate the forebrain are involved in sensory integration. Other sites that are strongly innervated by orexin fibers are the source of cholinergic projection of the brain stem and basal nuclei. The cholinergic projection of these areas is responsible for cortical activation, the result of which is the wakeful brain. The orexin projection is in charge of coordination of the activity of arousing cholinergic projections with the motor activity of the organism. The histaminergic cells of the posterior and other hypothalamic nuclei, which are engaged in the so-called forebrain wakefulness, are also under strict control of the orexin system [19]. The orexin projection is also in control of the dopaminergic neurons involved in reward processes and in the mechanism of wakefulness. Weaker projections innervate regions of the dorsal and ventral roots of the medulla oblongata, of motor neurons and limbic areas of the brainstem, as well as of the cerebral cortex [20] (Fig. 2). In the majority of cases, the orexin projection is stimulatory; this also refers to γ-aminobutyric acid-ergic (GABAergic) neurons of the pars reticulata substantia nigra or the septal nucleus innervating the hippocampus. Regarding the stimulatory action of the orexin projection, of paramount importance is the fact that it has no terminals of its own on neurons of specific thalamic pathways transmitting sensory information to the cerebral cortex [21]. This is further proof helping to classify this system as a nonspecific projection of the brain. The direct postsynaptic stimulatory action of orexin often depends on the effect of its simultaneous activation of other fibers that may have their terminals on the stimulated synapse. This happens, for example, in the case of serotonergic and noradrenergic cells that simultaneously receive direct orexin stimulation and indirect inhibition originating with GABAergic cells, also stimulated by orexin. Hence, the influence of the direct stimulatory action of orexin depends on indirect modulating effects and does not ever have to be of a stimulating nature. Such modulating action

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Fig. 2. The sagittal section through the rat brain shows the key projections of hypocretin (orexin) neurons (Hctr) from the lateral hypothalamus to the main components of the brain structures. See Fig. 1 caption for definitions of abbreviations. (After [20].)

that intensifies weak signals and inhibits strong ones is also a characteristic feature of nonspecific systems. The modulating effect of orexin projection on neuronal activity was also confirmed by the results of in vitro studies. The orexin-induced stimulation of orexin neurons in hypothalamic sections stems indirectly from the enhanced activity of glutaminergic cells adjacent to orexin ones but is not a direct effect on orexin cells [22]. The mechanism of inhibitory feedback can also be observed between the orexin system and the noradrenergic and serotoninergic ones. Orexin directly stimulates these two projections, which may attenuate its action on a feedback basis, via an inhibitory influence on the glutaminergic neurons present in the vicinity of the orexin projection. Also, the histaminergic projection originating with the protuberance of nuclei of the mammillary body exerts an inhibitory influence on orexin neurons via GABA release since histamine itself has a small effect on these neurons [22, 23]. Studies have shown that orexin release is endogenous and depends on the presence of the main generator of the mammalian biological clock: the SCNs of the hypothalamus [24]. The presence of OR-A and OR-B and OX-R1 receptors in the human retina seems particularly noteworthy [25] as this may suggest its modulatory role in the interaction among ganglionic cells of the retina, which transfer photic information to the SCN of the hypothalamus. Considering the presence of orexin OX-R1 receptors and OXcontaining fibers in the vicinity of the SCNs [25], it is proposed that this system may have a modulatory influence on the neuronal activity of the whole mechanism of mammalian biological clock. However, the unique role of orexin seems to be connected with its involvement—mediation of the transfer of nonphotic information to the intergeniculate leaflet (IGL) of the thalamus, the other (besides the SCN) extremely important neuronal element of the mechanism of the mammalian biological clock.

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INTERGENICULATE LEAFLET OF THE THALAMUS The IGL of the thalamus is the other (in addition to the SCN) structurally important element of the mechanism of mammalian biological clock. Anatomy The IGL belongs to the lateral geniculate nucleus (LGN) complex of the thalamus, an extremely important and interesting structure of the optic pathway. It is located between the dorsal lateral geniculate (DLG) and ventral lateral geniculate (VLG) parts of the lateral geniculate body. For a long time, the IGL was indistinguishable from the rest of the lateral geniculate body and was classified as in its ventral part. Only immunohistochemical and autoradiographic methods helped to demonstrate its complete anatomical identity and to define the limits of its occurrence and its connections with other brain structures [26, 27] (Fig. 3). The IGL is composed of small and medium-size multipolar interneurons with a dendritic zone that is limited to the area occupied by this structure, over which it differs anatomically from the remainder of the lateral geniculate body. This structure is clearly identifiable in rodents, whereas its homologs have so far been barely recognized in other species. In cats, such a homolog is the medial part of the ventral LGN, while in monkeys and humans it is the preginiculate nucleus [28]. The IGL receives strong innervation from retinal ganglionic cells of both eyes, branching off the RHT pathway running to the SCN. The projection to the contralaterally situated IGL is twice as large

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Fig. 3. The coronal section through the rat brain illustrating (left) the localization of the intergeniculate leaflet (IGL). A photomicrograph (right) of a rat brain coronal section across the geniculate complex with dark labeling of neuropeptide Y (NPY) immunoreactive neurons within the IGL area. DLG dorsal lateral geniculate nucleus, IGL intergeniculate leaflet, VLG ventral lateral geniculate nucleus.

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as that to the IGL situated ipsilaterally [29]. The above-mentioned leaflet stretches along the whole horizontal length of the LGN, occupying a section of about 2 mm in the rat and 2.2 mm in the hamster [27]. The majority of leaflet neurons reach the ventrolateral area of the SCN, bypassing—via the geniculohypothalamic tract (GHT)—neurons that form the RHT. The body of anatomic evidence showing the connection between the IGL and many other brain areas, especially those involved in the process of seeing, has been growing [30–34]. These relationships are generally bilateral and often reciprocal, being clear proof of a functional connection between the IGL and structures participating in the process of seeing, representing mainly the visuomotor function. This phenomenon was first investigated in 1994 by Morin [35], who confirmed it by additional proof, based mostly on his own anatomical studies published in an extremely interesting review, called most accurately, “The Circadian Visual System, 2005” [36]. It is noteworthy that there also exists a reciprocal bilateral connection between the two leaflets via the supraoptic commissure, which is limited to the IGL only, and therein to a population of neurons that do not project to the SCN. The role of this bilateral connection between the leaflets via the geniculogeniculate pathway is not entirely clear. It is thus assumed, although not yet conclusively proven, that both leaflets are reciprocally synchronized by means of this connection [37]. This synchronization seems to be particularly important with respect to the potential involvement of the IGL in the process of seeing, which requires synchronic activity on the part of all elements forming the visual system for the normal perception of an image. Possibly, the reciprocal connection between these two leaflets is another example of the integratory activity of brain. The Pharmacology of the IGL The total number of nerve cells forming the IGL ranges between 1,800 and 2,000 [27]. They are mostly GABAergic neurons that constitute a basic population of cells building up the neuronal mechanism of mammalian biological clock [38]. The pharmacological property by which the IGL differs from the remainder of the LGN is the presence of neuropeptide Y (NPY) (Fig. 3), which can be found mostly among GABAergic neurons projecting to the SCN [27, 39, 40]. Thus, NPY is regarded as the main transmitter of nonphotic information from the IGL to the SCN. Another neuropeptide present in the IGL is enkephalin (ENK), which in the majority of cases participates in the projection to the IGL situated opposite it. Also, neurotensin, which occurs in the vicinity of the majority of neurons with NPY, is a pharmacological marker of IGL neurons. Regarding the role of nonphotic information in the regulation of the mechanism of the biological clock, another very important IGL neuropeptide is relaxin 3. The extraordinarily rich nerve fibers containing this neuropeptide are present in numerous brain structures, yet above all in the IGL, being definitely less abundant in the SCN [41]. Relaxin probably participates in the modulation of behavioral patterns by adapting animals to environmental stressful conditions, which may constitute another nonphotic factor affecting the mechanism of the biological clock. The involvement of relaxin in the control of a visuomotor response has also been postulated [41], which in turn may confirm the engagement of the circadian system, above all the IGL, in visual processes.

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Chronobiology A vast body of anatomical evidence but above all the results of physiological and behavioral studies confirmed the involvement of the IGL in the regulation of biological rhythms. Electric stimulation of the IGL, or such nonphotic information as benzodiazepine treatment or novel wheel activity, produces a shift in the rhythm phase during the subjective day and a reduction in the expression of Period genes in the SCN [42], while light affects the circadian system during the subjective night, simultaneously bringing about an increase in the expression of the Period gene in the SCN. Electric lesion of both the IGLs always causes desynchronization of the rhythm of mouse locomotor activity [43]. The last effect may be explained by the lack of nonphotic synchronization in the circadian system after IGL lesion [44]. The observation that IGL lesion does not disturb the course of circadian rhythms in standard laboratory conditions under a specific permanent light regime (12 h of light/12 h of darkness) is of utmost importance. With limited access to other nonphotic stimuli, the strong light/darkness stimulus— always administered at the same time intervals—is a dominating signal that synchronizes rhythmic processes. However, it should be borne in mind that laboratory regimens are distant from the real conditions in which the majority of organisms live, above all humans. Therefore, the presence and significance of the structure that receives and integrates nonphotic information with photic information are of paramount importance. It is the IGL that fulfills this task. The Electrophysiology of the IGL The IGL neurons reveal an extremely interesting pattern of electric activity, characteristic only of this structure of the whole LGN complex. They generate action potentials in rhythmically repeated firing bursts with a constant interburst interval lasting several hours, defined as isoperiodic oscillations (Fig. 4). The mean time in which IGL cells change the level of their activity amounts to 124 ± 7 s [45]. Interestingly, such a pattern of activity is revealed by leaflet neurons only and is lacking in the dorsal and ventral parts of the LGN. On the other hand, these oscillations can be observed in the SCN of the hypothalamus [46, 47] and in the activity of cells of the pineal gland [48], to which the IGL has an additional projection in rats [49] and in

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Fig. 4. Firing rate histogram showing rhythmic slow bursting activity (isoperiodic, ultradian oscillation) of intergeniculate leaflet (IGL) neurons. Bin size 1 s.

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Mongolian gerbils [50]. The same pattern of cellular activity in the two most important structures (SCN and IGL) of the neuronal mechanism of mammalian biological clock permits an assumption that it constitutes a natural, extremely important basal rhythm characteristic of the work of not only these two structures, but also the whole mechanism of the mammalian biological clock. The absence of such oscillations in an in vitro preparation may suggest that they are of exogenous origin [51]. On the basis of the experiments conducted so far, in all probability it may be concluded that a photic signal from the retina is necessary for their development. Like the blockade of sodium conduction by means of TTX (tetrodotoxin citrate) administration to the eyeball, switching off the light inhibits the pattern of the oscillatory activity of IGL neurons [45, 52]. In this activity, mainly GABAergic neurons are involved; the blockade of their receptors leads to temporary disappearance of the oscillatory activity of the IGL [53]. It has also been ascertained that a reciprocal connection between both leaflets is not necessary for the occurrence of oscillatory activity in the opposite leaflet. The lesion or pharmacological blockade of one leaflet does not affect the oscillatory activity of the other [54]. Electric stimulation of the dorsal raphe nuclei—the main source of the serotoninergic projection to the IGL—causes a temporary decrease in the level of the oscillatory activity of neurons, while electric lesion of this projection results in a pronounced increase in this activity [55]. Our most recent studies with lesioned terminals of serotoninergic fibers in the IGL have confirmed the direct, distinct, modulatory role of the 5 hydroxytryptamine (5-HT) projection from the dorsal raphe nuclei in the oscillatory activity of IGL neurons [56]. A similar modulating effect is also observed in the case of two other nonspecific projections of the brain: cholinergic from the laterodorsal tegmental nuclei and noradrenergic from the locus coeruleus. Electrophysiological in vivo studies into the rat circadian system have shown an inhibitory effect of electric stimulation of the nonspecific projections of the brain stem on the potential induced in the IGL [57]. However, the influence of these two projections on the oscillatory activity of IGL neurons is not as apparent as that of the serotoninergic projection, which plays a dominating role in the mechanism of the mammalian biological clock. All the same, the crucial question about the role and significance of these short (rapid) oscillations for the functioning of the mammalian biological clock still remains unanswered. The physiological importance and connection of this ultradial rhythmicity of IGL and SCN neurons with long, commonly observed, and recorded circadian rhythms need to be elucidated. It is not easy to give an answer to this question at the present stage of knowledge. First, one should reflect on the issue whether this type of activity is limited exclusively to the mechanism of the biological clock. A similar activity, recorded by us lately in the pretectum (OPT) [58], permits us to assume that it may also be engaged in visual processes, in particular in visuomotor functions. This would provide us with another line of evidence for a close connection between the visual processes and regulation of biological rhythms and might account for the anatomical localization of the IGL in the structure involved in the process of seeing. However, the presence of oscillatory activity in SCN neurons—a structure engaged in the mechanism of the biological clock only—makes the involvement of oscillations of this type exclusively in visual processes questionable. One of the hypotheses (fairly universal, in my opinion) that explain the role of this rapid neuronal

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activity of the IGL predicts its possible engagement in the mechanism of release of NPY or orexins as well as in the pulsatory secretion of other neurohormones in the vicinity of the SCNs. Interestingly, only the eruptive activity of neurons is capable of facilitating the release of these macromolecular compounds from neuronal terminals. IGL as an Integrator of Photic and Nonphotic Information Both light and nonphotic nuclei are main synchronizers of the mechanism of the mammalian biological clock. Their interdependence and reciprocal connections are most clearly visible in environmental (extralaboratory) conditions. The neuronal element of the mechanism of the biological clock that integrates stimuli of either type is the IGL, while the orexin projection is probably a chemical transmitter of nonphotic information from nonspecific systems of the brain. The presence of terminals of OX-A fibers on the membrane of bodies of neurons with NPY in the IGL, in particular on those displaying gene c-Fos expression induced by locomotor activity (regarded as one of the more important nonphotic stimuli) [59–61], supports the above hypotheses. NPY, with neurons that richly innervate the SCN, is a transmitter of nonphotic information to this structure. The reciprocal integration of photic and nonphotic stimuli probably takes place on a molecular level via influence on the expression of Period genes in SCN neurons. In the light phase (in the daytime), a photic stimulus itself is not capable of changing the rhythm phase because the transcription of the gene Pre is already maximal in these light conditions. Only a nonphotic stimulus can lower gene Pre transcription in the light phase. Such lowering would enable a photic stimulus to exert an effect; it will try to inhibit the decrease in gene Period expression evoked by a nonphotic stimulus [62]. Orexin also modulates GLU release and stimulates GABAergic neurons (reviewed in [20]), the two main neurotransmitters of the biological clock. A direct effect of orexin on glutaminergic SCN cells seems unlikely as its receptors have been found in the vicinity of the SCN but not in the SCN itself. On the other hand, regulation of GLU secretion on the IGL level appears feasible on the assumption that GLU is a chemical transmitter of photic information to the IGL [63, 64]. All the above data give support to the hypothesis that the orexin system provides feedback between the mechanism of the mammalian biological clock determining circadian rhythms and the state of an organism’s activity. CONCLUSIONS The mechanism of the mammalian biological clock has two main oscillators: the SCN of the hypothalamus and the IGL of the thalamus. They form a closed, commonly working, synchronizing circadian system. The rhythmic work of this system is modulated by photic and nonphotic stimuli (Fig. 5). In humans, nonphotic stimuli are of greater importance for the system activity than is light. In contrast to light with a source, intensity of effect, and other parameters that can be precisely determined, nonphotic effects are difficult to precisely define as they belong to a group of stimuli that activate nonspecific systems of the brain, causing an arousal reaction that is transferred to the IGL, where photic stimuli are integrated with nonphotic ones.

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Fig. 5. The sagittal section through the rat brain shows the rough wiring between the main neuronal elements of the mammalian biological clock and the source of photic and nonphotic inputs. BF basal forebrain, GHT geniculohypothalamic tract, Hctr hypocretin, IGL intergeniculate leaflet, LC locus coeruleus, LDT laterodorsal tegmental nuleus, PPT penduculopontine tegmental nucleus, RHT retinohypothalamic tract, RN raphe nuclei, SCN suprachiasmatic nuclei, OPT optic tract.

A thorough knowledge of the mechanism of the circadian system synchronizing mammalian biological rhythms, precise identification of nonphotic effects, and knowledge of the neurobiological mechanism of their interactions are all of utmost importance for the understanding of many circadian dysfunctions that are the cause of the malfunctioning of the human organism. From a long-term perspective, the postulates presented—with the cooperation of pharmacologists—would permit possible interference in the work of the whole system, particularly improvement in the functioning of nonspecific cerebral systems that are a pivotal functional element of this system. Studies into the above-mentioned system would also stimulate development of new, not only pharmacological, but also behavioral therapeutic methods, permitting more efficient prevention of adverse states caused by functional disturbances to the circadian system that synchronizes biological rhythms in humans. This presents an interesting challenge not only to neurophysiologists, but also to psychologists, psychiatrists, and all the researchers who comprehensively analyze human organism. REFERENCES 1. Weaver, D. (1998) The suprachiasmatic nucleus: a 25-year retrospective. J. Biol. Rhythms 13, 100–112. 2. Moore, R. Y., Lenn, N. J. (1972) A retinohypothalamic projection in the rat. J. Comp. Neurol. 146, 1–14. 3. Rea, M. A. (1998) Photic entrainment of circadian rhythms in rodents. Chronobiol. Int. 5, 395–423. 4. Mrosovsky, N. (1996) Locomotor activity and non-photic influences on circadian clocks. Biol. Rev. Camb. Philos. Soc. 71, 343–372.

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5. Rietveld, W. J. (1996) General introduction to chronobiology. Braz. J. Med. Biol. Res. 29, 63–70. 6. Mistlberger, R. E., Skene, D. J. (2004) Social influences on mammalian circadian rhythms: animal and human studies. Biol. Rev. 79, 533–556. 7. Hasting, M. H., Duffield, G. E., Smith, E. J. D., Maywood, E. S., Ebling, F. J. P. (1998) Entrainment of the circadian system of mammals by nonphotic cues. Chronobiol. Int. 15, 425–445. 8. Moruzzi, G., Magoun, H. W. (1949) Brain stem reticular formation and activation of the EEG. Electroenceph. Clin. Neurophysiol. 1, 455–473. 9. Vrang, N., Mrosovsky, N., Mikkelsen, J. D. (2003) Afferent projections to the hamster intergeniculate leaflet demonstrated by retrograde and anterograde tracing. Brain Res. Bull. 59, 267–288. 10. Mistlberger, R. E., Antle, M. C., Glass J. D., Miller, J. D. (2000) Behavioral and serotonergic regulation of circadian rhythms. Biol. Rhythm Res. 31, 240–283. 11. Levitt, D.R., Teitelbaum, P. (1975) Somnolence, akinesia, and sensory activation of motivated behavior in the lateral hypothalamic syndrome. Proc. Natl. Acad. Sci. U. S. A. 72, 2819–2823. 12. De Lecea, L., Kilduff, T., Peyron, C., Gao, X. B., Foye, P. E., Danielson, P. E., Fukuhara, C., Battenberg, E. L., Gautvik, V. T., Bartlett, F. S. II., Frankel, W. N., van den Pol, A. N., Bloom, F. E., Gautvik, K. M., Sutcliffe, J. G. (1998) The hypocretins: Hypothalamus-specific peptides with neuroexcitatory activity. Proc. Natl. Acad. Sci. U. S. A. 95, 322–327. 13. Mileykovskiy, B. Y., Kiyashchenko, L. I., Siegel, J. M. (2005) Behavioral correlates of activity in identified hypocretin/orexin neurons. Neuron 46, 787–798. 14. Ripley, B., Overeem, S., Fujiki, N., Nevsimalova, S., Uchino, M., Yesavage, J., Di Monto, D., Dohi, K., Melberg, A., Lammers, G. J., et al. (2001) CSF hypocretin/orexin levels in narcolepsy and other neurological conditions. Neurology 57, 2253–2258. 15. Sakurai, T., Amemiya, A., Ishii, M., Matsuzaki, I., Chemelli, R. M., Tanaka, H., Williams, S. C., Richardson, J. A., Kozlowski, G. P., Wilson, S., Arch, J. R., Buckingham, R. E., Haynes, A. C., Carr, S. A., Annan, R. S., McNulty, D. E., Liu, W. S., Terrett, J. A., Elshourbagy, N. A., Bergsama, D. J., Yanagisawa, M. (1998) Orexins and orexin receptors: a family of hypothalamic neuropeptides and G protein-coupled receptors that regulate feeding behavior. Cell 92, 573–585. 16. Peyron, C., Tighe, D. K., van den Pol, A.N., de Lecea, L., Heller, H. C., Sutcliffe, J. G., Kilduff, T. S. (1998) Neurons containing hypocretin (orexin) project to multiple neuronal systems. J. Neurosci. 18, 9996–10015. 17. Ciriello, J., Rosas-Arellano, M. P., Solano-Flores, L. P., de Oliveira, C. V. (2003) Identification of neurons containing orexin-B (hypocretin-2) immunoreactivity in limbic structures. Brain Res. 967, 123–131. 18. Caillol, M., Aioun, J., Baly, C., Persuy, M. A., Salesse, R. (2003) Localization of orexin and their receptors in the rat olfactory system: possible modulation of olfactory perception by neuropeptide synthetized centrally or locally. Brain Res. 960, 48–61. 19. Nambu, T., Sakurai, T., Mizukami, K., Hosoya, Y., Yanagisawa, M., Goto, K. (1999) Distribution of orexin neurons in the adult rat brain. Brain Res. 827, 243–260. 20. Siegel, J.M. (2004) Hypocretin (orexin): role in normal behavior and neuropathology. Annu. Rev. Psychol. 55, 125–148. 21. Bayer, L., Eggermann, E., Saint-Mleux, B., Machard, D., Jones, B. E. (2002) Selective action of orexin (hypocretin) on nonspecific thalamocortical projection neurons. J. Neurosci. 22, 7835–7839.

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Lewandowski

22. Li, Y., Gao, X. B., Sakurai, T., van den Pol, A. N. (2002) Hypocretin/orexin excites hypocretin neurons via a local glutamate neurons – a potential mechanism for orchestrating the hypothalamic arousal system. Neuron 36, 1169–1181. 23. Yamanaka, A., Muraki, Y., Tsujino, N., Goto, K., Samurai, T. (2003) Regulation of orexin neurons by the monoaminergic and cholinergic systems. Biochem. Biophys. Res. Commun. 303, 120–129. 24. Zhang, S., Zeitzer, J. M., Yoshida, Y., Wisor, J. P., Nishino, S., Edgar, D. M., Mignot, E. (2004) Lesions of the suprachiasmatic nucleus eliminate the daily rhythm of hypocretin-1 release. Sleep 27, 619–627. 25. Savaskan, E., Müller-Spahn, F., Meier, F., Wirz-Justice, A., Meyer, P. (2004) Orexin and their receptors in the human retina. Pathobiology 71, 211–216. 26. Hickey, T. L., Spear, P. D. (1976) Retinogeniculate projections in hooded and albino rats. Exp. Brain Res. 24, 523–529. 27. Moore, R. Y., Card, J. P. (1994) Intergeniculate leaflet: an anatomically and functionally distinct subdivision of the lateral geniculate complex. J. Comp. Neurol. 344, 403–430. 28. Moore, R. Y. (1989) The geniculohypothalamic tract in monkey and man. Brain Res. 486, 190–194. 29. Morin, L. P., Blanchard, J. H., Provencio, I. (2003) Retinal ganglion cell projections to the hamster suprachiasmatic nucleus, intergeniculate leaflet, and visual midbrain: bifurcation and melanopsin immunoreactivity. J. Comp. Neurol. 465, 401–416. 30. Harrington, M. E. (1997) The ventral lateral geniculate nucleus and the intergeniculate leaflet: interrelated structures in the visual and circadian systems. Neurosci. Biobehav. Rev. 21, 705–727. 31. Moore, R. Y., Weis, R., Moga, M. M. (2000) Efferent projections of the intergeniculate leaflet and the ventral lateral geniculate nucleus in the rat. J. Comp. Neurol. 420, 398–418. 32. Morin, L., Blanchard J. H. (2001) Neuromodulator content of hamster intergeniculate leaflet neurons and their projection to the suprachiasmatic nucleus or visual midbrain. J. Comp. Neurol. 437, 79–90. 33. Vrang, N., Mrosovsky, N., Mikkelsen, J. D. (2003) Afferent projections to the hamster intergeniculate leaflet demonstrated by retrograde and anterograde tracing. Brain Res. Bull. 59, 267–288. 34. Horowitz, S. S., Blanchard, J. H., Morin, L.P. (2004) Intergeniculate leaflet and ventral lateral geniculate nucleus afferent connections: An anatomical substrate for functional input from the vestibule-visuomotor system. J. Comp. Neurol. 474, 227–245. 35. Morin, L. P. (1994) The circadian visual system. Brain Res. Rev. 67, 102–127. 36. Morin, L. P., Allen, C. N. (2006) The circadian visual system, 2005. Brain Res. Rev. 51, 1–60. 37. Mikkelsen, J. D. (1992) The organization of the crossed geniculogeniculate pathway of the rat: a Phaseolus vulgaris–leucoagglutinin study. Neuroscience 48, 953–962. 38. Moore, R. Y., Speh, J. C. (1993) GABA is the principal neurotransmitter of the circadian system. Neurosci Lett. 150, 112–116. 39. Morin, L. P., Blanchard, J. (1995) Organization of the hamster intergeniculate leaflet: NPY and ENK projections to the suprachiasmatic nucleus, intergeniculate leaflet and posterior limitans nucleus. Vis. Neurosci. 12, 57–67. 40. Janik, D., Mikkelsen, J. D., Mrosovsky, N. (1995) Cellular colocalization of Fos and neuropeptide Y in the intergeniculate leaflet after nonphotic phase-shifting events. Brain Res. 698, 137–145.

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41. Tanaka, M., Iijima, N., Miyamoto, Y., Fukusumi, S., Itoh, Y., Ozawa, H., Bata, Y. (2005) Neurons expressing relaxin 3/INSL 7 in the nucleus incertus respond to stress. Eur. J. Neurosci. 21, 1659–1670. 42. Yannielli, P., Harrington, M. E. (2004) Let there be “more” light: enhancement of light actions on the circadian system through non-photic pathways. Prog. Neurobiol. 74, 59–76. 43. Lewandowski, M. H., Usarek A. (2002) Effects of intergenculate leaflet lesions on circadian rhythms in the mouse. Behav. Brain Res. 128, 13–17. 44. Wickland, C., Turek, F. W. (1994) Lesions of the thalamic intergeniculate leaflet block activityinduced phase shifts in the circadian activity rhythm of the golden hamster. Brain Res. 660, 293–300. 45. Lewandowski, M. H., Błasiak, T., Domosławski, J., Wołkowska, A. (2000) Ultradian rhythmic neuronal oscillation in the intergeniculate leaflet. Neuroreport 11, 317–321. 46. Miller, J. D., Fuller, C. A. (1992) Isoperiodic neuronal activity in suprachiasmatic nucleus of the rat. Am. J. Physiol. 263, R51–R58. 47. Aggelopoulos, N. C., Meissl, H. (2000) Responses of neurones of the rat suprachiasmatic nucleus to retinal illumination under photic and scotopic conditions. J. Physiol. 523, 211–222. 48. Reuss, S. (1987) Electrical activity of the mammalian pineal gland. Pineal Res. Rev. 5, 153–189. 49. Mikkelsen, J. D., Moller, M. (1990) A direct neural projection from the intergeniculate leaflet of the lateral geniculate nucleus of the deep pineal gland of the rat, demonstrated with Phaseolus vulgaris leucoagglutinin. Brain Res. 520, 342–346. 50. Mikkelsen, J. D., Lozzi, B., Moller, M. (1991) Efferent projections from the lateral geniculate nucleus to the pineal complex of the Mongolian gerbil. Cell Tiss. Res. 264, 95–102. 51. Błasiak, A., Lewandowski, M. H. (2004) In vitro extracellular recording of spontaneous activity of the intergeniculate leaflet neurons. Brain Res. 1015, 82–86. 52. Lewandowski, M. H., Błasiak, T. (2004) Slow oscillation circuit of the intergeniculate leaflet. Acta Neurobiol. Exp. 64, 277–288. 53. Błasiak, T., Lewandowski, M. H. (2004) Blockade of GABAA receptors disrupts isoperiodic neuronal oscillations in the intergeniculate leaflet of the rat. Brain Res. 109, 82–87. 54. Lewandowski, M. H., Błasiak, T., Błasiak A. (2002) Are ultra-slow isoperiodic oscillations in rat intergeniculate leaflet neurons dependent on reciprocal connection with its contralaterally located counterpart? Neurosci. Lett. 330, 243–246. 55. Błasiak, T., Lewandowski, M. H. (2003) Dorsal raphe nucleus modulate isoperiodic neuronal activity in intergeniculate leaflet. Behav. Brain Res. 138, 179–185. 56. Błasiak, T., Siejka, S., Raison, S., Pevet, P., Lewandowski, M. H. (2006) Serotonergic inhibition of the slowly bursting cells in the intergeniculate leaflet of the rat. Eur. J. Neurosci. 24, 2769–2780. 57. Lewandowski, M. H., Błasiak, T. (1999) The effect of brainstem stimulation on the evoked potentials in the intergeniculate leaflet. Brain Res. 827, 139–142. 58. Szkudlarek, H., Lewandowski, M. H. (2005) IGL-like neuronal oscillations in the rat pretectum, similarities and discrepancies. Acta Neurobiol. Exp. 65, 327. 59. Nixon, J. P., Smale, L. (2004) Individual differences in wheel-running rhythms are related to temporal and spatial patterns of activation of orexin A and B cells in a diurnal rodent (Arvicanthis niloticus). Neuroscience 127, 25–34. 60. Nixon, J. P., Smale L. (2005) Orexin fibers from appositions with Fos expressing neuropeptide-Y cells in the grass rat intergeniculate leaflet. Brain Res. 1053, 33–37. 61. Thankachan, S., Rusak, B. (2005) Juxtacellular recording/labeling analysis of physiological and anatomical characteristics of rat intergeniculate leaflet neurons. J. Neurosci. 25, 9195–9204.

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62. Maywood, E. S., Mrosovsky, N. (2001) A molecular explanation of interactions between photic and non-photic circadian clock-resetting stimuli. Gene Expr. Patterns 1, 27–31. 63. Stamp, J. A., Piggins, H. D, Rusak, B., Semba, K. (1997) Distribution of ionotropic glutamate receptor subunit immunoreactivity in the suprachiasmatic nucleus and intergeniculate leaflet of the hamster. Brain Res. 756, 215–224. 64. Błasiak, A., Pękala, D., Lewandowski, M. H. (2007) The responsiveness of the rat intergeniculate leaflet neurons to glutamate agonists. J. Phsyiol. Pharmacol. 58, 669–681.

22 The Circadian Clock: Physiology, Genes, and Disease Michael C. Antle CONTENTS Introduction Entrainment Anatomy Clock Genes Human Implications Summary References

INTRODUCTION Most organisms on Earth live in cyclic environments created by the rotation of our planet around its axis. Daily environmental changes include geophysical variables such as light levels, temperature, barometric pressure, and relative humidity as well as biotic variables such as availability of food and presence of predators. To meet these predictable daily challenges, life has evolved a temporal organization regulated by an endogenous biological clock. This biological clock controls daily rhythms in both behavior and physiology. Even when housed in environments lacking temporal cues, organisms will continue to exhibit daily oscillations in their behavior and physiology, with periods close to, but not necessarily equal to, 24 h. As such, these endogenously generated oscillations are termed circadian rhythms, from the Latin circa “about” and dies “daily,” referring literally to any oscillation that repeats itself about daily. The circadian system has become a model system for understanding neural and molecular control of physiological and behavioral rhythms and has been implicated in a number of human sleep disorders. Circadian Rhythms in Physiology and Behavior The careful temporal coordination of physiological processes is essential for optimal functioning of our bodies [1]. For example, the physiological state required for seeking food (activation of the sympathetic nervous system) is quite different from that required for digesting food (activation of the parasympathetic nervous system), and the physiological From: Ophthalmology Research: Visual Transduction and Non-Visual Light Perception Edited by: J. Tombran-Tink and C. J. Barnstable © Humana Press, Totowa, NJ

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Fig. 1. A sample wheel-running record from a mouse. Each horizontal row represents the data from 1 day, with subsequent days plotted below previous days. The height of the black vertical lines in each row is proportional to the amount of wheel-running produced by the mouse in that 6-min bin. The animal is initially housed in a lighting cycle of 12 h light and 12 h dark. Note that the animal’s activity is initiated around the same time every day, just after the lights go out. Next, the animal is put into constant darkness. Note that while the animal maintains a daily pattern of activity, it now wakes up a little earlier each day, and it exhibits a free-running period of about 23.5 h. Finally, on the 12th day of darkness, the animal is exposed to a light pulse late in its subjective night (circle). This produces a phase advance in its activity onset, which can be visualized by fitting two regression lines to the activity onsets: one line on the days prior to the light pulse to predict the expected activity onset in the absence of a treatment (dark gray line) and another line on the days following the light pulse to determine the actual phase of the clock (white line). By comparing the horizontal distance between the two regression lines on the day after the light pulse, it is possible to quantify the magnitude of the phase shift. (Unpublished lab archive data; a color version is available on the accompanying CD.)

state of the body during sleep, including decreasing body temperature and secretion of melatonin and growth hormone, is distinct from each of the previous examples. Most physiological parameters exhibit daily oscillations. Prominent examples include body temperature, hormone secretions, and blood pressure. Behavioral parameters, such as locomotor activity (Fig. 1), also exhibit daily oscillations. The timing of sleep and wake onset and the timing of meals are also important examples of behaviors that demonstrate daily oscillations, but rhythms in human performance have also been noted. Reaction time is optimal in the late afternoon to early evening (4–8 p.m., and poorest in the early morning (4–8 a.m.) [2]. Search speed, reasoning speed, dexterity, and vigilance all exhibit poorest performance between 5 and 9 a.m., with a secondary trough occasionally observed in the early afternoon [3]. Performance on a memory task can also be influenced by a circadian rhythm [4]. Circadian Rhythms in Visual Function Visual performance and physiology exhibit circadian rhythms as well. In rats, there is a burst of rod outer segment disk shedding that coincides with the onset of light, but this rhythm persists for a number of days even in rats housed in constant darkness, suggesting that it is controlled by an endogenous clock rather than simply being a response to light exposure [5]. The axial length of the human eye changes by as much as 40 µm over the course of the day [6]. Intraocular pressure changes over the course of the day, with pressure high during the night, peaking near the end of the night, and lowest during the evening [7]. While this rhythm is largely affected by posture (higher when the person is

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supine) [7, 8], it may also be influenced by the flow of aqueous humor, which is lowest during the night (~1.3 µl/min) and highest during the day (~3 µl/min) [9]. The amplitude of the pupillary reflex can change over the course of the day [10]. Electroretinogram responses to stroboscopic flashes exhibit a circadian rhythm in the latency to the peak of the b-wave, with increasing latency throughout the night [11]. A circadian rhythm is observed in the electrooculogram produced by tracking moving objects, with maximal peak-to-peak amplitude in the late morning [11]. Performance on visual tasks also exhibits circadian rhythms in both acuity and sensitivity [11–14]. ENTRAINMENT While circadian oscillations in physiology and behavior are endogenously generated and regulated, they must be synchronized with the external geophysical and biotic cycles to serve an adaptive function. This is accomplished through entrainment, a process by which an external cue exerts period and phase control over a rhythm such that the endogenous period is adjusted to match that of the external environment (i.e., 24 h). The timing of the organism’s rhythms in relation to timing of events in the environment are also adjusted, such that nocturnal organisms are active at night, and diurnal organisms are active during the day. By controlling both the period and phase of the rhythms, entrainment ensures that physiological events occur at the proper times, and that it is possible to anticipate cyclic challenges. While many cues can provide time information to the circadian system and thus serve as zeitgebers (from the German zeit “time” and geber “giver”), not every cue has an impact on the circadian system in the same manner or to the same degree. Light is the dominant zeitgeber responsible for entrainment of circadian rhythmicity in most organisms. An intact retina and optic nerve are necessary for light to exert control over the mammalian circadian system [15–17] (although one claim to the contrary exists [18]). Light exposure early in the night will delay the phase of rhythms such that they occur later in subsequent cycles. Light exposure late in the night has the opposite effect and will advance the phase of rhythms [19] (Fig. 1). Light during the daytime has little effect on circadian rhythmicity. Through daily delay and advance adjustments, the endogenous circadian pacemaker has its period adjusted such that it equals that of the applied zeitgeber. Other cues serve as zeitgebers as well. These include dark pulses [20], exercise [21, 22], and sleep deprivation [23]. These nonphotic stimuli produce large phase advances during the midday and small delays late in the night but have little effect during the early night. A common feature of these zeitgebers is that they increase activity or wakefulness at a time of the day when the organism is generally inactive or asleep [23, 24]. While these phase shifts are independent of those produced by light exposure through the retina, there are interactions between the two zeitgeber systems. Exercise during a light pulse attenuates the resulting photic phase shift [25], possibly by inhibiting neurotransmitter release from retinal ganglion cells [25, 26]. Alternatively, housing animals in constant light for a number of days prior to a nonphotic pulse potentiates the resulting phase shift [27].

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ANATOMY The Suprachiasmatic Nucleus An accumulation of convergent evidence over the past 35 years has led to the conclusion that the master mammalian circadian pacemaker is located in the suprachiasmatic nucleus (SCN). This is a richly retinorecipient bilateral structure located at the base of the hypothalamus, adjacent to the third ventricle. SCN lesions abolish circadian rhythms in behavior and physiology [28, 29]. Transplants of fetal SCN to SCN-lesioned animals restores rhythmicity that exhibits the period characteristic of the donor, not the SCNlesioned host [30]. When cultured in vitro, individual SCN cells are able to maintain a circadian rhythm in their rate of spontaneous action potential generation [31]. While it appears that essentially every neuron in the SCN uses γ-aminobutyric acid (GABA) as a small molecule neurotransmitter [32], the SCN is actually a heterogeneous structure in terms of anatomy and function [33]. While it is difficult to delineate the myriad subregions of the SCN [34], the SCN has classically been subdivided into a ventrolateral “core” and a dorsomedial “shell.” These subdivisions were initially based on the heavily ventrolateral [35] pattern of retinal innervation and have been recapitulated with both phenotypic and functional heterogeneity [33]. Cells in the core receive retinal input, express the immediate early gene c-fos following a phase-shifting light pulse [36], and increase their firing rate during a light pulse [37] but do not exhibit an endogenous rhythm of electrical activity [38] or clock gene expression (see the section on clock genes). The core contains a number of different cell groups as defined by their peptidergic content, including cells containing vasoactive intestinal polypeptide (VIP), substance P, gastrin-releasing peptide (GRP), and calbindin (CalB) [33] (Fig. 2). In many cases, cells in the SCN core colocalize two or three of these peptides [39]. Much of what we know about the phenotypic heterogeneity of the SCN comes from examination of the Syrian hamster [33], and although there are some species differences, the peptidergic organization of the human SCN is quite similar to that observed in the hamster [40]. By contrast, cells in the SCN shell receive sparse retinal input [34], do not immediately respond with gene expression following a phase-shifting light pulse [41, 42], and do not change their firing rate during a light pulse [37] but do exhibit endogenous circadian rhythms in gene expression [41, 43]. The role of the SCN core appears to be to receive signals from the retina and relay them to the SCN shell. Retinal ganglion cells synapse onto cells containing CalB and GRP [44, 45]. These cell phenotypes occupy the region of the SCN with the heaviest retinal innervation [46]. VIP and GRP cells express c-fos after an animal has been exposed to a phase-shifting light pulse [47]. Application of VIP or GRP to the SCN in vivo and in vitro produces phase shifts of behavioral rhythms comparable to those observed following presentation of a light pulse (i.e., delays early in the night, advances late in the night [33, 48–52]). GRP receptors appear to occupy the shell region of the SCN [53]. These data indicate that certain cell phenotypes in the SCN core receive and respond to photic signals and produce signals that are able to mimic the effects of light. The core projects to cells in the shell but not vice versa [54]. Cells in the SCN core project heavily to regions outside the SCN as well, including the lateral septum, the ventrolateral preoptic area, the periventricular nucleus of the thalamus, the anterior portion of the periventricular

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Fig. 2. Sample photomicrographs depicting the location and the heterogeneity of the suprachiasmatic nucleus (SCN). Upper left A low-power nissl-stained coronal section through the anterior hypothalamus. The SCN on one side is outlined and is at the base of the third ventricle (3V), just dorsal to the optic chiasm (OX). The rest of the panels show the peptidergic phenotypes within the SCN detected using immunofluorescence. Depicted are calbindin-D28K (CalB), vasoactive intestinal polypeptide (VIP), vasopressin (VP), substance P (SP), and gastrin-releasing peptide (GRP). (Unpublished lab archive data; scale bars are 200 µm long.)

nucleus of the hypothalamus, the subperiventricular zone, the periaquaductal gray, and the intergeniculate leaflet (IGL) [54]. The SCN shell contains cells that are endogenously rhythmic. This conclusion is largely based on examination of clock gene expression patterns (see clock genes section), either using in situ hybridization or in vitro organotypic SCN cultures prepared from transgenic animals in which clock gene promoters drive the expression of either green fluorescent protein or luciferase. These cells do not respond immediately with gene expression [42] or increased firing rates [37] following a light pulse. This region is delineated by cells containing vasopressin, a peptide that is produced [43] and released [55] with a circadian rhythm. Cells in the shell project heavily to some of the same areas as the core as well as to the medial preoptic area, anterior hypothalamic area, the supraoptic nucleus, the dorsomedial nucleus of the hypothalamus, and the arcuate nucleus [54]. While the core and shell regions are useful as organizing concepts for understanding the heterogeneity of the SCN, they do not fully capture the complex organization of the SCN [34, 46]. By way of example, a third component of the SCN has been identified based on functional responses. This dorsolateral component of the SCN exhibits an antiphase oscillation in the phosphorylation of the extracellular signal-responsive kinases I/II (ERK). During the day, cells in the SCN shell contain the phosphorylated form of ERK (pERK). During the night, there is no pERK in the shell, but it is present in a distinct dorsolateral group of SCN cells [56]. Both of these patterns persist in timefree conditions, suggesting that they are not driven by the light–dark cycle but rather

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represent endogenously generated circadian rhythms. What is unique about the rhythmic ERK phosphorylation in the dorsolateral cells is that this oscillation requires innervation by the eye; enucleated animals no longer exhibit subjective night ERK phosphorylation in their dorsolateral SCN cells [56]. These cells are also involved in the photic phaseshifting pathway as microinjections of GRP selectively induce gene expression in these dorsolateral cells [49]. Inputs to the SCN The SCN receives input from a variety of sources, with about 40 brain areas providing monosynaptic input [57], expanding to around 100–140 brain areas when polysynaptic inputs are considered [58]. The three most prominent inputs to the SCN are the retinohypothalamic tract (RHT), the geniculohypothalamic tract (GHT), and serotonergic innervation from the midbrain raphe. The RHT originates in melanopsin-containing intrinsically photosensitive retinal ganglion cells [46]. The RHT projects to the ventrolateral SCN and uses glutamate as its primary neurotransmitter. The neuropeptides substance P and pituitary adenylate cyclaseactivating polypeptide have also been implicated in relaying information from retinal ganglion cells to the SCN [46]. NMDA (N-methyl-D-aspartate) produces photic phase shifts when applied to the SCN in vivo [59], and glutamate antagonists (both NMDA and non-NMDA antagonists) inhibit c-fos expression following a light pulse [60]. The second prominent source of SCN afferents is the IGL, which gives rise to the GHT. The IGL is a retinorecipient structure. It receives binocular input without apparent topographical organization [61]. The IGL also receives a variety of other inputs, including serotonergic input from the dorsal raphe [46]. IGL cells that innervate the SCN contain GABA and various neuropeptides, including neuropeptide Y (NPY), enkephalin, and neurotensin [46, 61]. The IGL appears to modulate both photic and nonphotic influences on the circadian system. Phase delays to light are enhanced in IGL-lesioned animals, while phase advances are attenuated [62]. Constant bright light normally lengthens a nocturnal animal’s endogenous period, an effect that is abolished in IGL-lesioned animals [62]. The IGL is also an integral component for nonphotic phase shifts. The IGL is the only source of NPY for the SCN, and injections of NPY to the SCN produce phase shifts similar to those produced by wheel confinement [63]. Lesions of the IGL prevent entrainment by enforced activity [64]. The third prominent region that inputs to the circadian system is the raphe complex. The median raphe innervates the SCN, while the dorsal raphe innervates the IGL. The prominent neurotransmitter of these structures is serotonin, although not all raphe projections are serotonergic [65]. The dorsal raphe is a retinorecipient structure [66, 67] and likely receives inputs from and sends outputs to the median raphe [68], thereby providing two more polysynaptic routes by which photic information can reach the SCN. In fact, some serotonergic drugs active at the serotonin 2C receptor have been reported to mimic photic phase shifts [69]. The serotonergic input from the raphe complex has been implicated in two major circadian functions. The first is modulation of nonphotic entrainment [70]. The evidence supporting serotonin’s role in nonphotic phase shifts is not as consistent as the evidence for NPY’s role in the same phenomenon [70]. Serotonin’s second role is to regulate photic input to the SCN [71]. Serotonin

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appears to attenuate neurotransmitter release from the RHT, but it may also be working postsynaptically [71]. Neurotoxic lesions of serotonergic input to the SCN can have a prominent effect on organization of behavioral rhythms. Animals have larger photic phase shifts, longer circadian periods under constant light, and an early phase of activity onset in a light–dark cycle [46, 72]. More extensive lesions of the serotonergic system can also affect the duration of the active phase [72]. Peripheral Oscillators A Clock in the Eye The discovery of oscillatory tissue outside the SCN has caused much excitement in the circadian rhythms field over the last decade. The first circadian oscillator to be discovered outside the SCN in mammals was in the retina [73]. The hamster retina tolerates culturing at 27°C, with the cultures being viable for a number of days. These retinal cultures release melatonin with a circadian rhythm, and this melatonin rhythm can still be entrained by a light–dark cycle [73]. Furthermore, the period of this rhythm is also affected by the tau mutation, resulting in a period of about 20 h [73]. A circadian rhythm in dopamine release has also been noted in the mouse retina and appears to depend on melatonin [74]. This circadian clock in the retina may mediate some of the circadian rhythms noted earlier in visual function and physiology to achieve optimal retinal functioning. Some of these functions may be controlled directly by the retinal clock, others may be controlled by one of the clock’s outputs (such as melatonin or dopamine), and still other functions may be controlled by the SCN. It is important to note that the human and primate retinas lack the final enzyme in the melatonin synthesis pathway (hydroxyindol O-methyltransferase) and therefore can only produce melatonin’s precursor, N-acetylserotonin, which may act as a signaling molecule in retinas lacking melatonin [75]. Melatonin appears to be released from photoreceptors in Xenopus and possibly the rat, while dopamine is released from amacrine and innerplexiform cells in mice [75]. Various genes driving the circadian clock (see clock genes section) are rhythmically expressed in a subset of retinal cells, specifically in both the inner nuclear layer and ganglion cell layer cells [76]. Amacrine cells containing dopamine, calbindin and/or calretinin, express clock genes, suggesting that these cells may contain autonomous circadian clocks [76]. In addition, bipolar cells and horizontal cells have been shown to express clock genes [77]. There is disagreement over whether cholinergic amacrine cells express clock genes [76, 77], but because only a small percentage of bipolar and cholinergic amacrine cells express clock genes, this may simply be a detection issue [77]. Oscillators Outside the Nervous System The identification of the genes underlying the intracellular circadian clock permitted the development of various tools to investigate clock gene expression, both within and beyond the SCN. Besides conventional approaches such as in situ hybridization, northern blot, and polymerase chain reaction (PCR), various novel approaches were developed

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using transgenic animals that used the promoters of different clock genes to drive the expression of reporters such as luciferase and green fluorescent protein. All of these approaches have led to the observation that clock genes are expressed outside the SCN [78] and indeed outside the brain [79]. Furthermore, clock gene expression is rhythmic in many of these tissues, including the liver, testes, and skeletal muscles [79]. It is possible that oscillations in gene expression in these peripheral tissues require the SCN as animals with SCN lesions do not exhibit circadian oscillations in gene expression in their liver [80]. This could be due to the loss of a daily organizing signal from the SCN, which maintains synchrony among the individual oscillating cells in these peripheral tissues. Cultured lung, skeletal muscle, liver, and SCN cells all exhibit circadian rhythms in gene expression as determined by real-time fluorescent reporters from a luciferase transgene driven by a promoter for one of the clock genes [81]. Even cultured fibroblasts can exhibit a circadian rhythm in gene expression if they are treated with a serum shock [82]. In all of these cultures, except for those of the SCN, oscillations decrease in amplitude and eventually extinguish after a number of cycles [81, 82]. This is believed to occur because, while each individual cell in these cultures continues to oscillate, synchrony among these oscillating cells is gradually lost [82–84]. Rhythmicity in these peripheral organs likely helps coordinate the diverse biochemical tasks carried out by these organs, such that incompatible biochemical reactions occur at different times, and metabolic reactions with high energetic demands are limited to only those times of the day when they are necessary [85]. CLOCK GENES The last decade or so has witnessed an explosion in our understanding of the molecular underpinnings of the mammalian circadian system, with the discovery of at least nine clock genes [1]. Our understanding of how these genes interact to yield the molecular circadian clock continues to evolve as recent findings have forced the field to reconsider the roles played by two of these genes [86, 87]. We now know that the circadian clock ticks at the level of single cells [31], and that this clock is a product of interlocked positive and negative transcription–translation feedback loops. While a genetic basis for circadian rhythmicity had long been suspected due to period mutants in fruit flies and Syrian hamsters as well as strain differences in period in mice, it was not until 1994 that the first clock gene was cloned in mammals. Clock was identified using a forward genetic approach by which colonies of mice were exposed to the mutagen ethylnitrosourea (ENU) and then screened for circadian rhythm abnormalities [88]. This gene is autosomal semidominant, causing lengthening of the circadian period from 23.3 h in wild-type animals, to 24.4 h in heterozygous animals, and to 27.2 h in homozygous animals. Homozygous Clock mutant animals quickly become arrhythmic in constant conditions. This gene codes for a basic helix-loop-helix (bHLH) transcription factor that contains a PAS dimerization domain (after the first three proteins found to contain such a domain: PER-ARNT-SIM; PER for period, ARNT for aryl hydrocarbon receptor nuclear translocator and SIM for single-minded protein). Clock dimerizes with BMAL1 (also known as MOP3) and together they act as positive regulators of genes that contain an Ebox (the DNA sequence CACGTG) in their promoter [89] (Fig. 3). Both of these genes

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Fig. 3. Schematic of the autoregulatory transcription–translation feedback loops that underlie the intracellular circadian clock. During the day, CLOCK/BMAL1 dimer drives the transcription of the Period, Cryptochrome, and Rev-erbα genes. The messenger RNA accumulates and is translated into proteins. PER and CRY proteins dimerize, and casein kinase 1ε phosphorylates PER proteins. In the late day to early night, the PER/CRY complex translocates to the nucleus (possibly with CK1ε), where it inhibits the activity of CLOCK and BMAL1 at E-boxes, thus terminating transcription. Transcription is reinitiated when the levels of PER and CRY proteins drop too low to continue to inhibit the activity of CLOCK and BMAL1. The auxiliary loop of rhythmic Rev-erbα production regulates the rhythmic production of BMAL1.

are highly conserved across phyla, and homologs have been found in a range of species, including Drosophila, in which BMAL1 is known as cycle. E-boxes regulate the expression of other clock genes as well as so-called clock-controlled genes such as vasopressin. It should be noted that while mutations of the clock gene impair the circadian clock, deletion of this gene has no effect on circadian rhythmicity [86]. The reasons for this are not

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currently known, but it is possible that another bHLH protein may dimerize with BMAL1 only when CLOCK protein, mutant or otherwise, is completely absent. The negative limb of the transcription–translation feedback loop is mediated by three Period and two Cryptochrome genes. All these genes have E-boxes in their promoters. The mammalian period gene Period1 (Per1) was first identified using a homology screen based on the sequence cloned for the Drosophila Period gene [90]. Two other homologs (Per2 and Per3) have since been discovered [79, 91, 92]. The peak levels of the messenger RNA (mRNA) for these genes are observed during the midday, with peak protein levels lagging behind by about 6 h. Mutations in either Per1 or Per2 alter various circadian properties. Per2 mutants have an abnormally short circadian period that degrades into arrhythmicity [93] and have attenuated phase delays to light [94]. Per1 mutants exhibit either shorter [94] or longer [95] periods and smaller phase advance [94]. When Per1 is constitutively expressed in transgenic rats, the period lengthens, and phase shifts to light are attenuated [96]. The Cryptochrome genes Cry1 and Cry2 were identified due to their homology with the blue light photoreceptor for the circadian systems of plants and flies. The peak levels of the mRNA for these genes are also observed during the midday. Disruption of these genes also results in period alterations, with longer periods observed when Cry2 is deleted and shorter periods observed when Cry1 is deleted [88, 97]. Animals lacking both Cry1 and Cry2 are completely arrhythmic when housed in constant darkness [88, 97]. The period and cryptochrome proteins are believed to dimerize and then translocate to the nucleus. Once in the nucleus, these complexes suppress the activity of CLOCK and BMAL1, thus terminating transcription of E-box-related genes. This suppression is largely attributed to the CRY proteins [98]. The leading hypothesis is that the CLOCK and BMAL1 are constitutively bound to the E-box, and that their presence leads to histone acetylation and thus chromatin remodeling that then permits RNA polymerase to bind to the promoter and initiate transcription of the gene. CRY is thought to inhibit the histone acetylation, which leads to chromatin changes that obscure the RNA polymerase promoter and prevent transcription of the gene [99]. As the Period and Cryptochrome genes are regulated by E-boxes, this negative-feedback loop has the effect of turning off the expression of these genes. The rhythmic expression in the SCN of the clock genes Per1 and Per2 as well as the clock-controlled gene vasopressin follows a specific spatiotemporal pattern and does not occur simultaneously in every cell [43, 100]. Expression is initiated early in the day in the dorsomedial SCN, spreads ventrally over the course of the day, and then recedes back to the dorsomedial SCN by the end of the day. Such rhythmic expression appears to be largely restricted to the SCN shell [41]. The circadian system also has a secondary loop that regulates the expression of BMAL1. The expression of the BMAL1 gene is regulated by a ROR (retinoid-related orphan receptor) promoter sequence. Two groups of proteins compete for the ROR sequence to regulate BMAL1 expression, namely, the ROR and Rev-erb families of proteins [101]. Specifically, the RORα isoform activate expression, while Rev-erbα isoform inhibits expression. Rev-erbα is itself regulated by an E-box and is expressed rhythmically, in phase with the Period and Cryptochrome genes and in antiphase to BMAL1 [101].

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One complete cycle of the transcription–translation feedback loop takes approximately 24 h. The rate of the these transcription–translation feedback loops that underlie the circadian clock are regulated by the casein kinase 1 family [102]. The first mammalian period mutation to be identified was the tau mutant hamster [103], a mutation subsequently mapped to the casein kinase 1ε (CK1ε) gene [102]. Hamsters that are heterozygous for this mutation have periods of approximately 22.2 h, while those that are homozygous for the mutation have periods of approximately 20.2 h [103]. CK1ε phosphorylates the period proteins, which alter their rates of translocation to the nucleus as well as their degradation rates. The tau mutation was initially thought to lead to decreased CK1ε-mediated phosphorylation of the PERIOD proteins, leading to premature peaks in PER1 and PER2 protein levels and in turn giving way to premature inhibition of CLOCK:BMAL1 activity [102]. This has been challenged by the observation that the tau mutation may actually increase phosphorylation of PER1 and PER2 and may lead to more rapid translocation to the nucleus or decreased degradation [87]. The expression of the Per1 and Per2 genes can also be regulated by input from the retina [79, 91, 92]. This induced expression is mediated by cAMP response element (CRE) elements in Per1 and Per2 promoters, which is regulated by the cyclic adenosine monophosphate (cAMP) response element-binding protein (CREB). Light pulses that phase shift the circadian clock phosphorylate CREB [104]. The initial induced expression of Per1 and Per2 occurs primarily in the SCN core and most prominently in the region of CalB cells [41]. Following a delay, some Per1 and Per2 expression is observed in the SCN shell. The pattern of this expression is related to the direction of the phase shift that is produced by the same light pulse. Only phase-delaying light leads to Per2 expression in the shell, while only phase-advancing light leads to Per1 expression in the shell [42]. Per1 and Per2 expression are essential components of the photic response; when their expression is prevented by microinjecting antisense oligonucleotides for these two genes into the SCN, phase shifts to light are attenuated or blocked [105]. The rest of the clock genes, including Per3, do not change their expression following exposure to phase-shifting light pulses. Manipulations that mimic light pulses also lead to increased expression of Period genes. The retina communicates with the SCN through glutamate release. NMDA microinjections to the SCN during the night produce phase shifts similar to those produced by light exposure at the same phases [59] and induce Per1 and Per2 expression [106]. Application of VIP to the SCN in vitro shifts the rhythm of electrical firing rates in a phase-dependent manner, such that early night application delays the phase of the peak firing rate, while application late in the night advances the peak to an earlier time. Such shifts in vitro are associated with increased Per1 and Per2 expression [107]. Microinjections of GRP to the SCN in vivo produce phase shifts in locomotor behavior comparable to those produced by light [52] and induce Per1 and Per2 expression in a discrete group of dorsolateral SCN cells [49]. While photic manipulations lead to an increased expression of Per1 and Per2, nonphotic manipulations have the opposite effect. Nonphotic manipulations, such as exercise and sleep deprivation, have their maximal effect at the same phase as peak rhythmic Per1 and Per2 expression (i.e., midday). These manipulations are associated with rapid downregulation of Per1 and Per2 [108]. In fact, decreasing Per1 expression

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directly by microinjecting Per1 antisense oligonucleotides to the SCN leads to nonphotic phase shifts [109]. HUMAN IMPLICATIONS People differ in their preferred sleep patterns. Some people are “night owls” who like to stay up late and sleep in if possible, while other people are “morning larks” who prefer to go to sleep early and wake up early. Such patterns are often developmental; the night owl pattern is frequently observed during adolescence, while the morning lark pattern is more common in the elderly [110]. This developmental pattern has been attributed to a change in the period of the circadian clock over the life span [110]. In some people, these patterns can be quite extreme, leading to the clinical diagnoses of either advanced or delayed sleep phase syndrome (ASPS or DSPS, respectively). In ASPS, the patient typically initiates sleep around 7:30 p.m., waking up around 4:30 a.m., while in DSPS, sleep often starts between 2 and 6 a.m. In some cases, these syndromes appear to be heritable. Given that mutations in clock genes can alter properties of the circadian clock in animals, is it possible that these human sleep disorders may be due to polymorphisms in clock genes? A mutation in the human Per2 gene has been identified in one group with heritable ASPS [111]. This mutation affects the binding site on the Per2 protein for CK1ε, the enzyme that is altered in the tau mutant hamster [102]. Like humans with ASPS, the tau mutant hamster also has an advanced circadian phase when entrained [103]. A mutation in the related enzyme CK1δ in humans has also been associated with ASPS [112]. DSPS can also be heritable and is associated with a polymorphism in the human Per3 gene [113] in some cases and a missense mutation in the CK1ε gene in other cases [114]. Both of these disorders can be managed to a limited degree with timed light exposure [115]. Early-morning bright light can help with DSPS, while ASPS can be treated with evening light. SUMMARY The circadian system is an ideal model for understanding neural regulation of complex behavioral and physiological phenomena at the level of the tissue, the cell, and the molecule. The study of the circadian system is relevant not only to understanding daily changes in behavior and physiology but also for optimizing therapeutic regimens that may have a circadian rhythm (so-called chronopharmacology). Furthermore, the study of the circadian system is important for understanding human diseases that may exhibit daily fluctuations in symptoms, or that may result from an aberration in the circadian system itself, as is observed in ASPS and DSPS. REFERENCES 1. Hastings, M. H., Reddy, A. B., Maywood, E. S. (2003). A clockwork web: circadian timing in brain and periphery, in health and disease. Nat Rev Neurosci 4, 649–661. 2. Wright, K. P., Jr., Hull, J. T., Czeisler, C. A. (2002). Relationship between alertness, performance, and body temperature in humans. Am J Physiol Regul Integr Comp Physiol 283, R1370–R1377.

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3. Monk, T., Buysse, D., Reynolds, C., III, Berga, S., Jarrett, D., Begley, A. M. Y., Kupfer, D. (1997). Circadian rhythms in human performance and mood under constant conditions. J Sleep Res 6, 9–18. 4. May, C. P., Hasher, L., Stoltzfus, E. R. (1993). Optimal time of day and the magnitude of age differences in memory. Psychol Sci 4, 326–330. 5. LaVail, M. M. (1976). Rod outer segment disk shedding in rat retina: relationship to cyclic lighting. Science 194, 1071–1074. 6. Stone, R. A., Quinn, G. E., Francis, E. L., Ying, G. S., Flitcroft, D. I., Parekh, P., Brown, J., Orlow, J., Schmid, G. (2004). Diurnal axial length fluctuations in human eyes. Invest Ophthalmol Vis Sci 45, 63–70. 7. Liu, J. H., Sit, A. J., Weinreb, R. N. (2005). Variation of 24-hour intraocular pressure in healthy individuals: right eye versus left eye. Ophthalmology 112, 1670–1675. 8. Hara, T., Hara, T., Tsuru, T. (2006). Increase of peak intraocular pressure during sleep in reproduced diurnal changes by posture. Arch Ophthalmol 124, 165–168. 9. Brubaker, R. F. (1991). Flow of aqueous humor in humans [the Friedenwald lecture]. Invest Ophthalmol Vis Sci 32, 3145–3166. 10. Fosnaugh, J. S., Bunker, E. B., Pickworth, W. B. (1992). Daily variation and effects of ambient light and circadian factors on the human light reflex. Methods Find Exp Clin Pharmacol 14, 545–553. 11. Tuunainen, A., Kripke, D. F., Cress, A. C., Youngstedt, S. D. (2001). Retinal circadian rhythms in humans. Chronobiol Int 18, 957–971. 12. Bassi, C. J., Powers, M. K. (1986). Daily fluctuations in the detectability of dim lights by humans. Physiol Behav 38, 871–877. 13. Tassi, P., Pellerin, N., Moessinger, M., Eschenlauer, R., Muzet, A. (2000). Variation of visual detection over the 24-hour period in humans. Chronobiol Int 17, 795–805. 14. Tassi, P., Pellerin, N., Moessinger, M., Hoeft, A., Muzet, A. (2000). Visual resolution in humans fluctuates over the 24 h period. Chronobiol Int 17, 187–195. 15. Eastman, C. I., Martin, S. K., Hebert, M. (2000). Failure of extraocular light to facilitate circadian rhythm reentrainment in humans. Chronobiol Int 17, 807–826. 16. Wright, K. P., Jr., Czeisler, C. A. (2002). Absence of circadian phase resetting in response to bright light behind the knees. Science 297, 571. 17. Yamazaki, S., Goto, M., Menaker, M. (1999). No evidence for extraocular photoreceptors in the circadian system of the Syrian hamster. J Biol Rhythm 14, 197–201. 18. Campbell, S. S., Murphy, P. J. (1998). Extraocular circadian phototransduction in humans. Science 279, 396–399. 19. Daan, S., Pittendrigh, C. S. (1976). A functional analysis of circadian pacemakers in nocturnal rodents. II. The variability of phase response curves. J Comp Physiol 106, 253–266. 20. Boulos, Z., Rusak, B. (1982). Circadian phase response curves for dark pulses in the hamster. J Comp Physiol 146, 411–417. 21. Buxton, O. M., Lee, C. W., L’Hermite-Baleriaux, M., Turek, F. W., Van Cauter, E. (2003). Exercise elicits phase shifts and acute alterations of melatonin that vary with circadian phase. Am J Physiol Regul Integr Comp Physiol 284, R714–R724. 22. Van Reeth, O., Sturis, J., Byrne, M. M., Blackman, J. D., L’Hermite-Baleriaux, M., Leproult, R., Oliner, C., Refetoff, S., Turek, F. W., Van Cauter, E. (1994). Nocturnal exercise phase delays circadian rhythms of melatonin and thyrotropin secretion in normal men. Am J Physiol 266, E964–E974. 23. Antle, M. C., Mistlberger, R. E. (2000). Circadian clock resetting by sleep deprivation without exercise in the Syrian hamster. J Neurosci 20, 9326–9332.

494

Antle

24. Reebs, S. G., Lavery, R. J., Mrosovsky, N. (1989). Running activity mediates the phaseadvancing effects of dark pulses on hamster circadian rhythms. J Comp Physiol A 165, 811–818. 25. Mistlberger, R. E., Antle, M. C. (1998). Behavioral inhibition of light-induced circadian phase resetting is phase and serotonin dependent. Brain Res 786, 31–38. 26. Rea, M. A., Glass, J. D., Colwell, C. S. (1994). Serotonin modulates photic responses in the hamster suprachiasmatic nuclei. J Neurosci 14, 3635–3642. 27. Mistlberger, R. E., Belcourt, J., Antle, M. C. (2002). Circadian clock resetting by sleep deprivation without exercise in Syrian hamsters: Dark pulses revisited. J Biol Rhythm 17, 227–237. 28. Stephan, F. K., Zucker, I. (1972). Circadian rhythms in drinking behavior and locomotor activity of rats are eliminated by hypothalamic lesions. Proc Natl Acad Sci U S A 69, 1583– 1586. 29. Moore, R. Y., Eichler, V. B. (1972). Loss of a circadian adrenal corticosterone rhythm following suprachiasmatic lesions in the rat. Brain Res 42, 201–206. 30. Ralph, M. R., Foster, R. G., Davis, F. C., Menaker, M. (1990). Transplanted suprachiasmatic nucleus determines circadian period. Science 247, 975–978. 31. Welsh, D. K., Logothetis, D. E., Meister, M., Reppert, S. M. (1995). Individual neurons dissociated from rat suprachiasmatic nucleus express independently phased circadian firing rhythms. Neuron 14, 697–706. 32. Moore, R. Y., Speh, J. C., Leak, R. K. (2002). Suprachiasmatic nucleus organization. Cell Tissue Res 309, 89–98. 33. Antle, M. C., Silver, R. (2005). Orchestrating time: arrangements of the brain circadian clock. Trends Neurosci 28, 145–151. 34. Morin, L. P., Shivers, K. Y., Blanchard, J. H., Muscat, L. (2006). Complex organization of mouse and rat suprachiasmatic nucleus. Neuroscience 137, 1285–1297. 35. Moore, R. Y. (1996). Entrainment pathways and the functional organization of the circadian system. Prog Brain Res 111, 103–119. 36. Silver, R., Romero, M. T., Besmer, H. R., Leak, R., Nunez, J. M., LeSauter, J. (1996). Calbindin-D28K cells in the hamster SCN express light-induced Fos. Neuroreport 7, 1224– 1228. 37. Meijer, J. H., Groos, G. A., Rusak, B. (1986). Luminance coding in a circadian pacemaker: the suprachiasmatic nucleus of the rat and the hamster. Brain Res 382, 109–118. 38. Jobst, E. E., Allen, C. N. (2002). Calbindin neurons in the hamster suprachiasmatic nucleus do not exhibit a circadian variation in spontaneous firing rate. Eur J Neurosci 16, 2469– 2474. 39. LeSauter, J., Kriegsfeld, L. J., Hon, J., Silver, R. (2002). Calbindin-D(28K) cells selectively contact intra-SCN neurons. Neuroscience 111, 575–585. 40. Mai, J. K., Kedziora, O., Teckhaus, L., Sofroniew, M. V. (1991). Evidence for subdivisions in the human suprachiasmatic nucleus. J Comp Neurol 305, 508–525. 41. Hamada, T., LeSauter, J., Venuti, J. M., Silver, R. (2001). Expression of Period genes: rhythmic and nonrhythmic compartments of the suprachiasmatic nucleus pacemaker. J Neurosci 21, 7742–7750. 42. Yan, L., Silver, R. (2002). Differential induction and localization of mPer1 and mPer2 during advancing and delaying phase shifts. Eur J Neurosci 16, 1531–1540. 43. Hamada, T., Antle, M. C., Silver, R. (2004). Temporal and spatial expression patterns of canonical clock genes and clock-controlled genes in the suprachiasmatic nucleus. Eur J Neurosci 19, 1741–1748.

Circadian Rhythms

495

44. Bryant, D. N., LeSauter, J., Silver, R., Romero, M. T. (2000). Retinal innervation of calbindin-D28K cells in the hamster suprachiasmatic nucleus: ultrastructural characterization. J Biol Rhythm 15, 103–111. 45. Tanaka, M., Hayashi, S., Tamada, Y., Ikeda, T., Hisa, Y., Takamatsu, T., Ibata, Y. (1997). Direct retinal projections to GRP neurons in the suprachiasmatic nucleus of the rat. Neuroreport 8, 2187–2191. 46. Morin, L. P., Allen, C. N. (2006). The circadian visual system, 2005. Brain Res Rev 51, 1–60. 47. Romijn, H. J., Sluiter, A. A., Pool, C. W., Wortel, J., Buijs, R. M. (1996). Differences in colocalization between Fos and PHI, GRP, VIP and VP in neurons of the rat suprachiasmatic nucleus after a light stimulus during the phase delay versus the phase advance period of the night. J Comp Neurol 372, 1–8. 48. Albers, H. E., Liou, S. Y., Stopa, E. G., Zoeller, R. T. (1991). Interaction of colocalized neuropeptides: functional significance in the circadian timing system. J Neurosci 11, 846–51. 49. Antle, M. C., Kriegsfeld, L. J., Silver, R. (2005). Signaling within the master clock of the brain: localized activation of mitogen-activated protein kinase by gastrin-releasing peptide. J Neurosci 25, 2447–2454. 50. McArthur, A. J., Coogan, A. N., Ajpru, S., Sugden, D., Biello, S. M., Piggins, H. D. (2000). Gastrin-releasing peptide phase-shifts suprachiasmatic nuclei neuronal rhythms in vitro. J Neurosci 20, 5496–5502. 51. Meyer-Spasche, A., Piggins, H. D. (2004). Vasoactive intestinal polypeptide phaseadvances the rat suprachiasmatic nuclei circadian pacemaker in vitro via protein kinase A and mitogen-activated protein kinase. Neurosci Lett 358, 91–94. 52. Piggins, H. D., Antle, M. C., Rusak, B. (1995). Neuropeptides phase shift the mammalian circadian pacemaker. J Neurosci 15, 5612–5622. 53. Karatsoreos, I. N., Romeo, R. D., McEwen, B. S., Silver, R. (2006). Diurnal regulation of the gastrin-releasing peptide receptor in the mouse circadian clock. Eur J Neurosci 23, 1047–1053. 54. Kriegsfeld, L. J., Leak, R. K., Yackulic, C. B., LeSauter, J., Silver, R. (2004). Organization of suprachiasmatic nucleus projections in Syrian hamsters (Mesocricetus auratus): an anterograde and retrograde analysis. J Comp Neurol 468, 361–379. 55. Kalsbeek, A., Buijs, R. M., Engelmann, M., Wotjak, C. T., Landgraf, R. (1995). In vivo measurement of a diurnal variation in vasopressin release in the rat suprachiasmatic nucleus. Brain Res 682, 75–82. 56. Lee, H. S., Nelms, J. L., Nguyen, M., Silver, R., Lehman, M. N. (2003). The eye is necessary for a circadian rhythm in the suprachiasmatic nucleus. Nat Neurosci 6, 111–112. 57. Krout, K. E., Kawano, J., Mettenleiter, T. C., Loewy, A. D. (2002). CNS inputs to the suprachiasmatic nucleus of the rat. Neuroscience 110, 73–92. 58. Pickard, G. E., Smeraski, C. A., Tomlinson, C. C., Banfield, B. W., Kaufman, J., Wilcox, C. L., Enquist, L. W., Sollars, P. J. (2002). Intravitreal injection of the attenuated pseudorabies virus PRV Bartha results in infection of the hamster suprachiasmatic nucleus only by retrograde transsynaptic transport via autonomic circuits. J Neurosci 22, 2701–2710. 59. Mintz, E. M., Marvel, C. L., Gillespie, C. F., Price, K. M., Albers, H. E. (1999). Activation of NMDA receptors in the suprachiasmatic nucleus produces light-like phase shifts of the circadian clock in vivo. J Neurosci 19, 5124–5130. 60. Abe, H., Rusak, B., Robertson, H. A. (1992). NMDA and non-NMDA receptor antagonists inhibit photic induction of Fos protein in the hamster suprachiasmatic nucleus. Brain Res Bull 28, 831–835.

496

Antle

61. Harrington, M. E. (1997). The ventral lateral geniculate nucleus and the intergeniculate leaflet: interrelated structures in the visual and circadian systems. Neurosci Biobehav Rev 21, 705–727. 62. Pickard, G. E., Ralph, M. R., Menaker, M. (1987). The intergeniculate leaflet partially mediates effects of light on circadian rhythms. J Biol Rhythm 2, 35–56. 63. Huhman, K. L., Albers, H. E. (1994). Neuropeptide Y microinjected into the suprachiasmatic region phase shifts circadian rhythms in constant darkness. Peptides 15, 1475–1478. 64. Marchant, E. G., Watson, N. V., Mistlberger, R. E. (1997). Both neuropeptide Y and serotonin are necessary for entrainment of circadian rhythms in mice by daily treadmill running schedules. J Neurosci 17, 7974–7987. 65. Stamp, J. A., Semba, K. (1995). Extent of colocalization of serotonin and GABA in the neurons of the rat raphe nuclei. Brain Res 677, 39–49. 66. Fite, K. V., Janusonis, S., Foote, W., Bengston, L. (1999). Retinal afferents to the dorsal raphe nucleus in rats and Mongolian gerbils. J Comp Neurol 414, 469–484. 67. Shen, H., Semba, K. (1994). A direct retinal projection to the dorsal raphe nucleus in the rat. Brain Res 635, 159–168. 68. Glass, J. D., Grossman, G. H., Farnbauch, L., DiNardo, L. (2003). Midbrain raphe modulation of nonphotic circadian clock resetting and 5-HT release in the mammalian suprachiasmatic nucleus. J Neurosci 23, 7451–7460. 69. Kennaway, D. J., Moyer, R. W. (1998). Serotonin 5-HT2C agonists mimic the effect of light pulses on circadian rhythms. Brain Res 806, 257–270. 70. Mistlberger, R. E., Antle, M. C., Glass, J. D., Miller, J. D. (2000). Behavioral and serotonergic regulation of circadian rhythms. Biol Rhythm Res 31, 240–283. 71. Rea, M. A., Pickard, G. G. E. (2000). Serotonergic modulation of photic entrainment in the Syrian hamster. Biol Rhythm Res 31, 284–314. 72. Meyer-Bernstein, E. L., Blanchard, J. H., Morin, L. P. (1997). The serotonergic projection from the median raphe nucleus to the suprachiasmatic nucleus modulates activity phase onset, but not other circadian rhythm parameters. Brain Res 755, 112–120. 73. Tosini, G., Menaker, M. (1996). Circadian rhythms in cultured mammalian retina. Science 272, 419–421. 74. Doyle, S. E., Grace, M. S., McIvor, W., Menaker, M. (2002). Circadian rhythms of dopamine in mouse retina: the role of melatonin. Vis Neurosci 19, 593–601. 75. Iuvone, P. M., Tosini, G., Pozdeyev, N., Haque, R., Klein, D. C., Chaurasia, S. S. (2005). Circadian clocks, clock networks, arylalkylamine N-acetyltransferase, and melatonin in the retina. Prog Retin Eye Res 24, 433–456. 76. Witkovsky, P., Veisenberger, E., LeSauter, J., Yan, L., Johnson, M., Zhang, D. Q., McMahon, D., Silver, R. (2003). Cellular location and circadian rhythm of expression of the biological clock gene Period 1 in the mouse retina. J Neurosci 23, 7670–7676. 77. Ruan, G. X., Zhang, D. Q., Zhou, T., Yamazaki, S., McMahon, D. G. (2006). Circadian organization of the mammalian retina. Proc Natl Acad Sci U S A 103, 9703–9708. 78. Sun, Z. S., Albrecht, U., Zhuchenko, O., Bailey, J., Eichele, G., Lee, C. C. (1997). RIGUI, a putative mammalian ortholog of the Drosophila period gene. Cell 90, 1003–1011. 79. Zylka, M. J., Shearman, L. P., Weaver, D. R., Reppert, S. M. (1998). Three period homologs in mammals: Differential light responses in the suprachiasmatic circadian clock and oscillating transcripts outside of brain. Neuron 20, 1103–1110. 80. Akhtar, R. A., Reddy, A. B., Maywood, E. S., Clayton, J. D., King, V. M., Smith, A. G., Gant, T. W., Hastings, M. H., Kyriacou, C. P. (2002). Circadian cycling of the mouse liver transcriptome, as revealed by cDNA microarray, is driven by the suprachiasmatic nucleus. Curr Biol 12, 540–550.

Circadian Rhythms

497

81. Yamazaki, S., Numano, R., Abe, M., Hida, A., Takahashi, R., Ueda, M., Block, G. D., Sakaki, Y., Menaker, M., Tei, H. (2000). Resetting central and peripheral circadian oscillators in transgenic rats. Science 288, 682–685. 82. Balsalobre, A., Damiola, F., Schibler, U. (1998). A serum shock induces circadian gene expression in mammalian tissue culture cells. Cell 93, 929–937. 83. Welsh, D. K., Yoo, S. H., Liu, A. C., Takahashi, J. S., Kay, S. A. (2004). Bioluminescence imaging of individual fibroblasts reveals persistent, independently phased circadian rhythms of clock gene expression. Curr Biol 14, 2289–2295. 84. Antle, M. C., Foley, D. K., Foley, N. C., Silver, R. (2003). Gates and oscillators: a network model of the brain clock. J Biol Rhythm 18, 339–350. 85. Schibler, U., Naef, F. (2005). Cellular oscillators: rhythmic gene expression and metabolism. Curr Opin Cell Biol 17, 223–229. 86. Debruyne, J. P., Noton, E., Lambert, C. M., Maywood, E. S., Weaver, D. R., Reppert, S. M. (2006). A clock shock: mouse CLOCK is not required for circadian oscillator function. Neuron 50, 465–477. 87. Gallego, M., Eide, E. J., Woolf, M. F., Virshup, D. M., Forger, D. B. (2006). An opposite role for tau in circadian rhythms revealed by mathematical modeling. Proc Natl Acad Sci U S A 103, 10618–10623. 88. Vitaterna, M. H., King, D. P., Chang, A. M., Kornhauser, J. M., Lowrey, P. L., McDonald, J. D., Dove, W. F., Pinto, L. H., Turek, F. W., Takahashi, J. S. (1994). Mutagenesis and mapping of a mouse gene, Clock, essential for circadian behavior. Science 264, 719–725. 89. Hogenesch, J. B., Gu, Y. Z., Jain, S., Bradfield, C. A. (1998). The basic-helix-loop-helixPAS orphan MOP3 forms transcriptionally active complexes with circadian and hypoxia factors. Proc Natl Acad Sci U S A 95, 5474–5479. 90. Tei, H., Okamura, H., Shigeyoshi, Y., Fukuhara, C., Ozawa, R., Hirose, M., Sakaki, Y. (1997). Circadian oscillation of a mammalian homologue of the Drosophila period gene. Nature 389, 512–516. 91. Albrecht, U., Sun, Z. S., Eichele, G., Lee, C. C. (1997). A differential response of two putative mammalian circadian regulators, mPer1 and mPer2, to light. Cell 91, 1055–1064. 92. Shearman, L. P., Zylka, M. J., Weaver, D. R., Kolakowski, L. F., Jr., Reppert, S. M. (1997). Two period homologs: circadian expression and photic regulation in the suprachiasmatic nuclei. Neuron 19, 1261–1269. 93. Zheng, B., Larkin, D. W., Albrecht, U., Sun, Z. S., Sage, M., Eichele, G., Lee, C. C., Bradley, A. (1999). The mPer2 gene encodes a functional component of the mammalian circadian clock. Nature 400, 169–173. 94. Albrecht, U., Zheng, B., Larkin, D., Sun, Z. S., Lee, C. C. (2001). mPer1 and mPer2 are essential for normal resetting of the circadian clock. J Biol Rhythm 16, 100–104. 95. Zheng, B., Albrecht, U., Kaasik, K., Sage, M., Lu, W., Vaishnav, S., Li, Q., Sun, Z. S., Eichele, G., Bradley, A., Lee, C. C. (2001). Nonredundant roles of the mPer1 and mPer2 genes in the mammalian circadian clock. Cell 105, 683–694. 96. Numano, R., Yamazaki, S., Umeda, N., Samura, T., Sujino, M., Takahashi, R., Ueda, M., Mori, A., Yamada, K., Sakaki, Y., Inouye, S. T., Menaker, M., Tei, H. (2006). Constitutive expression of the Period1 gene impairs behavioral and molecular circadian rhythms. Proc Natl Acad Sci U S A 103, 3716–3721. 97. van der Horst, G. T., Muijtjens, M., Kobayashi, K., Takano, R., Kanno, S., Takao, M., de Wit, J., Verkerk, A., Eker, A. P., van Leenen, D., Buijs, R., Bootsma, D., Hoeijmakers, J. H., Yasui, A. (1999). Mammalian Cry1 and Cry2 are essential for maintenance of circadian rhythms. Nature 398, 627–630.

498

Antle

98. Kume, K., Zylka, M. J., Sriram, S., Shearman, L. P., Weaver, D. R., Jin, X., Maywood, E. S., Hastings, M. H., Reppert, S. M. (1999). mCRY1 and mCRY2 are essential components of the negative limb of the circadian clock feedback loop. Cell 98, 193–205. 99. Ripperger, J. A., Schibler, U. (2006). Rhythmic CLOCK-BMAL1 binding to multiple E-box motifs drives circadian Dbp transcription and chromatin transitions. Nat Genet 38, 369–374. 100. Yamaguchi, S., Isejima, H., Matsuo, T., Okura, R., Yagita, K., Kobayashi, M., Okamura, H. (2003). Synchronization of cellular clocks in the suprachiasmatic nucleus. Science 302, 1408–1412. 101. Sato, T. K., Panda, S., Miraglia, L. J., Reyes, T. M., Rudic, R. D., McNamara, P., Naik, K. A., FitzGerald, G. A., Kay, S. A., Hogenesch, J. B. (2004). A functional genomics strategy reveals Rora as a component of the mammalian circadian clock. Neuron 43, 527–537. 102. Lowrey, P. L., Shimomura, K., Antoch, M. P., Yamazaki, S., Zemenides, P. D., Ralph, M. R., Menaker, M., Takahashi, J. S. (2000). Positional syntenic cloning and functional characterization of the mammalian circadian mutation tau. Science 288, 483–492. 103. Ralph, M. R., Menaker, M. (1988). A mutation of the circadian system in golden hamsters. Science 241, 1225–1227. 104. Ginty, D. D., Kornhauser, J. M., Thompson, M. A., Bading, H., Mayo, K. E., Takahashi, J. S., Greenberg, M. E. (1993). Regulation of CREB phosphorylation in the suprachiasmatic nucleus by light and a circadian clock. Science 260, 238–241. 105. Wakamatsu, H., Takahashi, S., Moriya, T., Inouye, S. T., Okamura, H., Akiyama, M., Shibata, S. (2001). Additive effect of mPer1 and mPer2 antisense oligonucleotides on lightinduced phase shift. Neuroreport 12, 127–131. 106. Moriya, T., Horikawa, K., Akiyama, M., Shibata, S. (2000). Correlative association between N-methyl-D-aspartate receptor-mediated expression of period genes in the suprachiasmatic nucleus and phase shifts in behavior with photic entrainment of clock in hamsters. Mol Pharmacol 58, 1554–1562. 107. Nielsen, H. S., Hannibal, J., Fahrenkrug, J. (2002). Vasoactive intestinal polypeptide induces per1 and per2 gene expression in the rat suprachiasmatic nucleus late at night. Eur J Neurosci 15, 570–574. 108. Maywood, E. S., Mrosovsky, N., Field, M. D., Hastings, M. H. (1999). Rapid down-regulation of mammalian period genes during behavioral resetting of the circadian clock. Proc Natl Acad Sci U S A 96, 15211–15216. 109. Hamada, T., Antle, M. C., Silver, R. (2004). The role of Period1 in non-photic resetting of the hamster circadian pacemaker in the suprachiasmatic nucleus. Neurosci Lett 362, 87–90. 110. Myers, B. L., Badia, P. (1995). Changes in circadian rhythms and sleep quality with aging: mechanisms and interventions. Neurosci Biobehav Rev 19, 553–571. 111. Toh, K. L., Jones, C. R., He, Y., Eide, E. J., Hinz, W. A., Virshup, D. M., Ptacek, L. J., Fu, Y. H. (2001). An hPer2 phosphorylation site mutation in familial advanced sleep phase syndrome. Science 291, 1040–1043. 112. Xu, Y., Padiath, Q. S., Shapiro, R. E., Jones, C. R., Wu, S. C., Saigoh, N., Saigoh, K., Ptacek, L. J., Fu, Y. H. (2005). Functional consequences of a CKIdelta mutation causing familial advanced sleep phase syndrome. Nature 434, 640–644. 113. Ebisawa, T., Uchiyama, M., Kajimura, N., Mishima, K., Kamei, Y., Katoh, M., Watanabe, T., Sekimoto, M., Shibui, K., Kim, K., Kudo, Y., Ozeki, Y., Sugishita, M., Toyoshima, R., Inoue, Y., Yamada, N., Nagase, T., Ozaki, N., Ohara, O., Ishida, N., Okawa, M., Takahashi, K.,

Circadian Rhythms

499

Yamauchi, T. (2001). Association of structural polymorphisms in the human period3 gene with delayed sleep phase syndrome. EMBO Rep 2, 342–346. 114. Takano, A., Uchiyama, M., Kajimura, N., Mishima, K., Inoue, Y., Kamei, Y., Kitajima, T., Shibui, K., Katoh, M., Watanabe, T., Hashimotodani, Y., Nakajima, T., Ozeki, Y., Hori, T., Yamada, N., Toyoshima, R., Ozaki, N., Okawa, M., Nagai, K., Takahashi, K., Isojima, Y., Yamauchi, T., Ebisawa, T. (2004). A missense variation in human casein kinase I epsilon gene that induces functional alteration and shows an inverse association with circadian rhythm sleep disorders. Neuropsychopharmacology 29, 1901–1909. 115. Reid, K. J., Burgess, H. J. (2005). Circadian rhythm sleep disorders. Prim Care 32, 449–473.

Index

10° viewing fields 336 11-cis retinal 69, 88, 92, 146, 175 11-cis retinol dehydrogenase (11cRDH) 396, 435 2° brightness-matching luminous efficiency function 338 2° photopic viewing conditions 336 2° viewing fields 335 4-hydroxy-2-nonenal 428 5 hydroxytryptamine (5-HT) 474 A A2E see N-retinylidene-N-retinylethanolamine A2E/iso-A2E 426 A2E-phosphatidyletholamine (A2-PE) 393, 397 Achromatopsia 237, 243, 245 Adenosine diphosphate (ADP) ribosylation 152 Adenosine 204 Advanced glycation endproducts (AGE) 412 Advanced sleep phase syndrome (ASPS) 492 Age-related macular degeneration (AMD) 81, 395 Akt/P13K 47 All-trans retinal (ATR) 69, 92, 146, 175, 424 All-trans retinol 69, 88 Amacrine cells 6, 22, 36,39,293 AMPA receptors 293 AMPA 289 Amygdala 469 Anchoring INAD 274 APB (L-2-amino-4 phosphononbutyric acid) 377 Apoprotein E (apoE) 396 Aqueous humor 5 Arachidoneate 424 Arousal reaction 467

ARPE-19 79 Arr2 275 Arrestin 175 Ascending reticular-activating system (ARAS) 467 Astigmatism 5 Atonal 37 ATP-binding cassette transporter (ABCR) 234, 279, 393, 396, 431 B Best’s disease 423 Best’s vitelliform macular dystrophy 437 Bestrophin-1 437 bHLH genes 39 Bipolar cells 6, 10, 22, 36, 39, 207,360 Bipolar-to-ganglion cell synapses 292 Blind spot 9 Blood-retinal barrier 68 Blue-cone monochromacy 319 Blue-green spectrum 344 BMAL1 22, 488 Bone morphogenetic protein (BMP) 21 Bruch’s membrane 68 bZIP 262 C Ca2+/calmodulin-dependent protein kinase II (CaMKII) 132, 236 Cadherin-based adherens junctions 68 Calbindin (CalB) 202, 484 Calcium 144, 197, 225, 227 Calcium-binding protein (CaBP) 202 Calcium-induced calcium release (CICR) 207 Calcium-recoverin 147 Calmodulin 143 Calpains 210 Calpastatin 211

501

502 cAMP-dependent protein kinase (PKA) 142 Casein kinase 1ε (CK1ε) 491 Caspase 1 211 Caspase cascade 210 Catabolite activator protein (CAP) 231 Catalase 414 CBF-1 21 CBF3 21 Ccr-2 439 CD36 78 Cellular retinal-binding protein (CRALBP) 435 CEP-protein 447 Channel-exchanger complex 279 Chondroitinase ABC 71 Choriocapillaris 395 Choroid 96 Chromatic channels 331, 344 Chx10 41, 107, 112 Ciliary muscles 3 Ciliary neurotrophic factor (CNTF) 47 Cilium 226 Circadian rhythms 74 CK1δ 492 CLF/CLC 48 CLOCK 490 CNTF receptor (CNTFRα) 47 Cobblestone 406 Color blind 289 Color vision 251, 307, 365 Color-matching functions (CMFs) 309, 313 Color-sensitive opsin 252, 255, 262 COMP1 108, 112 Complement factor H (CFH) 396 Complete achromatopsia 319 Cone photoreceptor 6, 36, 261, 268, 353 dystrophy 237 fundamentals 311 monochromacies 308, 319 photopigment 365 pigment polymorphisms 343 pigment spectra 355 signal pathways 359 spectral sensitivities 307, 310, 313 threshold 329 Cone-based vision 361 Cone rod homeobox gene (Crx) 40, 49, 90, 103, 261 Cone-rod break 319

Index Cone-rod dystrophy (CORD2) 98 Congenital color vision deficiencies 314 Congenital stationary night blindness (CSNB) 185, 377, 382, Connecting cilium 144 Cornea 3 Cortical 395 CRALBP 449 CRE 491 CREB 491 Critical flicker fusion CFF 332 Crx see Cone rod homeobox Crx-binding element (CBE) 104 Cry2 490 Cryptochrome genes Cry1 490 c-Src 132 Cyclic adenosine monophosphate (cAMP) 141 Cyclic adenosine monophosphate (cAMP) 181 Cyclic guanosine monophosphate (cGMP) 141, 225, 269 Cyclic guanosine monophosphate -gated cation channels 132, 141, 225, 243, 269, 277, 367 Cyclic guanosine monophosphate (cGMP) phosphodiesterase 130, 141 Cyclic guanosine monophosphate-dependent protein kinases (PKGs) 141 CYP26 21 D Dark current 227 D-cis diltiazem 212 Deuteranomaly 316 Deuteranopia 315 Diacylglycerol 143, 424 Dichromacy 308 Directional sensitivity 344 Directional-selective ganglion cells 293, 300 Discs-large protein (hDlg) 271 Disk shedding 75 dl-buthionine-(S,R)-sulfoximine 447 Discs large gene (Dlg) 271 Docosahexaenoic acid (DHA) 45, 424, 427 Dopamine 204, 468 Dorsal lateral geniculate (DLG) 471 Dorsal rim area (DRA) 253 Drusen 404, 415

Index Deep sleep phase syndrome (DSPS) 492 DTL (Dawson-Trick-Litzkow) 376 E Enhanced S-cone syndrome (ECSC) 44 Electroretinogram (ERG) 319, 357 Ellipsoid 144 ELOVL4 437 Endoplasmic reticulum (ER) 199, 236 Enhanced S-cone syndrome (ESCS) 43 Enkephalin (ENK) 472, 486 Entrainment 483 Eph A3 21 Ephrin B1 21 Ephrin B2 21 Epidermal growth factor (EGF) 46, 257 Experimental autoimmune uveitis (EAU) 88 External limiting membrane (ELM) 68 Extracellular receptor kinase (ERK) 236 Eye-specific protein kinase C (ePKC) 271 Ezrin 70 F FAK (focal adhesion kinase) 79, 81 FGF8 21 Fibroblast growth factor (FGF) 133, 212 Focal oscillatory potentials (fOPs) 375 Fodrin 210 Forebrain wakefulness 469 Fourier-transform infrared (FTIR) 180 Fovea 6, 17, 333, 391 Foveal and parafoveal cones 333 development 20, 26 hypoplasia 27 location 20 pit formation 27 pit 24, 28 Foveola 391 Foxd1 21 Foxg1 21 Frizzled transmembrane receptors 46 Frizzled-4 46 Fundus autofluorescence 415 Fundus flavimaculatis 415 Fundus 404 Furano-A2E 447

503 G G protein activation 277 G protein molecule 307 G protein-coupled receptor kinase 1 (GRK1) 127 coupled receptors 251 coupled signaling pathways 267 receptor-protein rhodopsin 307 GABA (γ-aminobutyric acid) 204, 377 GABA aminotransferase 386 GABAA receptor 295 GABAC receptor-mediated presynaptic inhibition 294 GABAC receptors 295 GABAergic lateral inhibition 295, 298 GABAergic L-IPSCs 295 Ganglion cell excitatory responses 292 Ganglion cell layer (GCL) 4, 17, 107 Ganglion cells 6, 18, 90 GAP (guanosine triphosphatase (GTPase) accelerating protein 131 Gap junctional connections 212, 299 GARP (glutamine acid-rich protein) 229, 245, 279 Gastrin-releasing peptide (GRP) 484 Gate chloride channels 298 GC-activating proteins (GCAPs) 147 Geniculohypothalamic tract (GHT) 486 GH6 21 Glial cells 13 Glutamate 204, 475 receptors 292 release 291 transporters 204, 292 Glutathione peroxidase (GPX) 414 Glycine 47, 298 Glycine receptor α2 (GlyRa2) 47 Glycinergic amacrine cells 299 Goldman-Favre syndrome 43 Gp130 47 Gq protein 269 GRK7 130 Guanosine diphosphate (GDP) 227 Guanosine triphosphatase (GTPase) 146, 278 Guanosine triphosphate (GTP) 142, 227 Guanylate cyclase (GC) 141, 209, 228 Guanylate cyclase-activating protein (GCAP) 209, 228

504 H Helix-loop-helix (bHLH) 488 Heterochromatic brightness matching (HBM) 332 Heterochromatic flicker photometry (HFP) 332 Heterochromatic modulation photometry (HMP) 332 High-gain phosphorylation 129 Horizontal cells 6, 22, 36, 90 Human color-sensitive opsins 260 Hybrid LM or ML cone photopigment opsin genes 316 Hydrophobic interference 186 Hydroxyl radical (OH•) 413 Hyperopia 5 Hypocretin (Hctr) 468 Hypoxia 208 I Intergeniculate leaflet (IGL) 471 INAD (inactivation no afterpotential D) 268, 271, Inflammatory phagocytosis 78 Inner nuclear layer (INL) 4, 17, 39, 40, 49 Inner plexiform layer (IPL) 4, 287,289 Inner segment (IS) 4, 7, 68, 198 Inositol triphosphate (IP3) 199, 269 Insulin 204, 259 Insulin-like growth factor I (IGF-I) 132, 235 Intergeniculate leaflet (IGL) 470, 485 Interleukin 6 (IL-6) 47 Interleukin 8 (IL-8) 447 Interphotoreceptor matrix (IPM) 67, 70, 88 Interphotoreceptor retinoid-binding protein (IRBP) 69, 73, 87, 90, 94, 100, 110, 114 Interphotoreceptor space (IPS) 87 Intradiskal domain 183 Intrinsically photosensitive ganglion cells 300 Iodopsin 74 Ionotropic GABA receptors 295 IP3 see Inositol triphosphate Iris 5 Ischemia 211 J JAK 47, 49

Index K Krüppel-like factor 15 (KLF15) 106 L L/M cones 43 L-and M-cone spectral sensitivities 312 Lateral Geniculate nucleus (LGN) 471 Lateral paths 11 Lawrence-Moon-Biedl syndrome 415 L-cis Diltiazem 212 L-cone see long-wavelength sensitive cone Lens pigment 313 Lens 3 LIF 47 LIFRβ 47 Light adaptation 144, 159 Light/dark cycle 437 Lipid peroxidation 409, 412, 443 Lipid-derived aldehydes 443 Lipofuscin 393, 421 Lipophilic granules 421 L-IPSCs 295 Locus coeruleus 468 Locus control region (LCR) 257 Long-wavelength sensitive cone (L cone) 18, 312, 380 L-opsin gene 369 Love spot 253 LRAT 448 LRP6 46 L-type calcium channels 291 Luminance pathway 332 Luminous efficiency functions 330 Lumirhodopsin 177 Lutein 18 M M cones see Medium-wavelength sensitive cone M/L pigments 359 Macula leutea 17 Macula 18 Macular pigment 314, 341 Maf transcription factor family 41 M-cone opsin 49, 356 MDB 332 Medium-wavelength sensitive cone (M-cone) 18, 355, 380

Index Meis1 113 Melanin 422 Melanocortin MC1R receptor 183 Melanolipofuscin 422 MERTK 73, 80, 427 Mesopic (rod-cone) luminous efficiency functions 339 Metarhodopsin 127, 146, 307 mfERGs (multifocal electroretinogram) 377, 385 mfOPs 375 mGluR receptors 206 Microphthalmia transcription factor 41, 72 Microvillar membrane 275 Midget ganglion cells 299 Mitf see microphthalmia transcription factor Mitochondria 144, 199 Mitochondrial Ca2+ uniporters 202 Mitochondrial DNA (mtDNA) 393, 409 Mitogen-activated protein kinase (MAPK) 47, 132 MOK2 107 Monochromacies 308, 319 Monochromats 310 Monocyte chemoattractant protein 1 (MCP-1) 438, 439, 447 Mouse cone pigments 355 Müller cells 13, 22, 36, 48, 68, 107 Multifocal oscillatory porential (fOP) 377, 382 Mutagen ethylnitrosourea (ENU) 488 Myopia 5 Myosin NINAC (neither inactivation no afterpotential C) 271 N N-(4-hydroxyphenyl) retinamide (HPR) 396, 434 Na/Ca-K exchanger 279 Na+/K+,Ca2+ exchanger 227 Narrow-field amacrine cell 293 Neofunctionalization 96 Nerve fiber layer 4, 89 Neural retina leucine zipper (Nrl) 23, 41, 50, 90, 102, 114, 262 Neural-cell adhesion molecule (N-CAM) 72 Neuraminidase 71 Neuro D 37, 39, 49 Neuropeptide Y (NPY) 486

505 Neuropoeitin 48 Neurotensin 486 Nicotinamide adenine dinucleotide phosphate (NADPH) 433 Night blindness 134 Nitric oxide (NO) 204, 206, 228,429 Nitric oxide synthase (NOS) 413, 429 NMDA (N-methyl-D-aspartate) 486 NMDA receptors 293 NO see nitric oxide Nocturnal 483 Nonproliferative diabetic retinopathy (NPDR) 385 Noradrenaline 468 Neuropeptide Y (NPY) 472, 475 Nr2E 49 Nr2E3 23, 43, 45, 262 N-retinylamides 448 N-retinylidene phosphatidylethanolamine (NRPE) 432, 441 N-Retinylidene-N-retinylethanolamine (A2E) 393, 409 NRF2 450 Nrl see Neural retina leucine zipper Nuclear DNA (nDNA) 409 Nuclear receptors 43 Nucleoside 3′,5′-cyclic monophosphates (cNMPs) 229 O Ocular retardation mouse mutant 107 OFF bipolar cell 11, 18 OFF sublaminae 291 Oguchi disease 129 Ommatidia 252 ON and OFF bipolar cells 289 ON and OFF sublaminae 291 ON bipolar cells 11 ON sublaminae 291 ONL 41, 101, 105,107 OPL 288 Opsin 7,47,77,81,251,427 Opsin degradation 175 Opsin kinase see Rhodopsin kinase Optic disk 9 Optokinetic nystagmus (OKN) 362 Orexin (OX) 468 Organic anion-transporting protein (OATP) 446

506 Otx1 41 Otx2 39, 103,113 Otx5b 41 Outer nuclear layer (ONL) 4, 40, 88 Outer plexiform layer (OPL) 4, 287 Outer segment see Photoreceptor outer segment P p561ck 132 Parallel bipolar cell-signaling pathways 299 Parallel ganglion cell output pathways 299 Parvalbumin 202 Pax2 21, 113 Pax6 22, 27, 41, 261 Pbx 113 PDE6 see Cyclic guanosine monophosphate (cGMP) phosphodiesterase PDZ domains 271 PER1 491 PER2 490 Per3 490 Peripheral vision 171 Peripherin 234, 279 Periphery 333 Permeability transition pore 209 Peroxy-A2E 447 Peroxynitrites (ONOO•) 413 Phagocytosis 75. 77, 427 Phosducin 132 Phosphatidylcholine 424 Phosphatidylethanolamine 424 Phosphatidylinositol 424 Phosphatidylserine 424 Phosphodiesterase (PDE) see also Cyclic guanosine monophosphate (cGMP) phosphodiesterase 8, 227, 269 Phosphodiesterase β subunit 209 Phosphoinositol signaling pathway 269 Phosphoinositol-bis-phosphate (PIP2) 269 Photoisomerization 69, 145 Photophobia 237, 243 Photopic brightness-matching function 340 Photopic luminous efficiency functions 334 Photopic vision 334, 395 Photopigment optical density 314, 332, 342 Photopigment variability 316 Photoreception 5, 268

Index Photoreceptor 4, 68, 73, 81, 89, 197, 225, 243, 252 Photoreceptor differentiation 48 Photoreceptor guanylate cyclase (GC) 130 Photoreceptor ion channels 269 Photoreceptor layer 6 Photoreceptor membrane 278 Photoreceptor- Müller cell junctions 68 Photoreceptor outer segment 4, 7, 67, 73, 77, 80, 92, 197, 268, 424 Photoreceptor retinol dehydrogenase (prRDH) 431 Photoreceptor rim protein see ATP-binding cassette transporter (ABCR) Photoresponse 144 Phototransduction 7, 69, 74, 267, 278, 307 Phototransduction proteins 275 Pigment dispersion 415 Pingelapese blindness 245 Plasma membrane Ca2+ adenosine triphosphatase (ATPase) transporters (PMCAs) 202 PLCβ 271 Pleiotrophin 48 PNA 354 Poly ADP ribose polymerase (PARP) 210 POS see Photoreceptor outer segment Postsynaptic excitatory postsynaptic current (EPSC) 208 Prenyl-binding protein (PrBP/δ) 158 Prep1 113 Preretinal screening 332 Presbyopia 4 Presynaptic inhibition 294 Prokineticin 1 439 Protan and deutan defects 315 Protanomaly 316 Protanopia 315 Protein kinase A (PKA) 132 Protein kinase C (PKC) 130, 236 Protein oxidation (protein carbonyl formation) 409 Protein phosphatase 2A (PP2A) 133 Protein phosphatase A (PPA) 175 Protein tyrosine phosphatases (PTPs) 235 prRDH 432 PSD95 271 Pupil 5

Index Q QRX homeobox gene 41 R Radial glia 212 Retinaldehyde dehydrogenase (RALDH) 21 Rapid-eye movement (REM) 469 Rax-L 41 RBP3 112 Rd7 43 Reactive oxygen species (ROS) 209, 403 Receptor tyrosine kinase Mer (MerTK) 79 Recoverin 48, 129, 147, 202 Red cone monochromats 319 Red-green deficiencies 315 Refractive index 3 Relevant light specification (photometry) 330 Ret1/PCE I 114 Retinal blood vessels 26 Retinal capillary network 18 Retinal chromophore 145 Retinal detachment 208 Retinal G protein-coupled receptor (RGR) 435 Retinal ganglion cells 12, 22, 36 Retinal histogenesis 36 Retinal pigment epithelium (RPE) 4, 7, 39, 46, 67, 77, 87, 92, 96, 175 Retinal progenitor cells (RPCs) 36, 37, 44, 47, 212 Retinal β-ionone ring 183 Retinitis pigmentosa (RP) 81, 129, 182, 208, 237, 367, 415, 423 Retinohypothalamic tract (RHT) 466, 471, 486 Retinoic acid receptors (RARs) 43, 45 Retinoic acid 45, 88 Retinoid 45, 69, 87, 91,143 Retinoid binding proteins 88 Retinoid orphan receptor beta (RORβ) 43 Retinoid related orphan receptor β (RORβ) 359 Retinoid X receptors (RXRs) 43 Retinol 425 Retinol acyl transferase (LRAT) 434 Retinol dehydrogenase (RDH) 432 RDH5 435, 449 RDH11 435 RDH12 433

507 Retinol-binding protein (RBP) 434 Retinyl palmitate 425 Rh5 253 Rh6 253 Rhabdomere 252, 268 Rhodopsin kinase (RK) 125, 158, 175 Rhodopsin 8, 37, 42, 46,69, 102, 126, 171, 228, 252, 268, 367 Rhythm 466 Rod dark adaptation 129 Rod monochromacy 320 Rod monochromats 308 Rod photoreceptor 6, 22, 36, 90, 261, 353, 392 development 35, 40, 46, 49 spectral sensitivity 333 threshold 329 Rod-based signals 360 Rod-cone luminous efficiency functions 333 Rod-cone self-cancellation 339 Rod-free zone 22 ROR (retinoid-related orphan receptor) 490 RORβ 43 RPE65 72, 396, 449 RPE-J 79 Rx 41, 101 Rx-L 41 RXR 43, 45, 113 Ryanodine 199 S S/UV pigments 359 Sarcoplasmic-endoplasmic reticulum Ca2+ ATPase (SERCA) 199 Sclera 3 Suprachiasmatic Nucleus (SCN) 470, 484 S-cone see Short-wavelength-sensitive cone S-cone monochromats 312 S-cone ON bipolars 360 S-cone opsin 45, 49 S-cone spectral sensitivity 310, 312 Scotopic luminous efficiency 332 Scotopic vision 171, 334, 395 Scotopic-to-photopic transition 339 Serotonin 486 Short-wavelength-sensitive cones (S-cones) 18, 42, 44, 310, 358 Signal-responsive kinases I/II (ERK) 485

508 Singlet oxygen 443 Site-directed cysteine mutagenesis 180 SOHo-1 21 Somatostatin 204 Spalt gene complex (sal) 257 Spatial and temporal sensitivity 364 Specific matching or detection tasks 331 Spectral distribution 333 Spectral luminous efficiency 331 Spectral sensitivity 330, 363 Spillover transmission 294 Stargardt’s disease 423 STAT3 48 STDG3 437 Steric hindrance 186 Subfunction partitioning 96 Sulfhydryl modification 180 Superoxide anion (O2•) 413 Superoxide dismutase (SOD) 410 Superoxide radical 443 Suppressor of cytokine signaling 3 (SOCS3) 49 Suprachiasmatic nuclei (SCNs) 466 Sustained (X) fashion 299 Sustained and transient response stratification 291 SWS1 family 356 Synaptic ribbons 292 T Taurine 47 Tbx5 21 TDH (3,7,11-trimethyldodeca-2,6,10-trienoic acid hexadecylamide) 435 TDT (13,17.21-trimethyldocosa-12,16, 20-trien-11-one) 435 Tert-butyl hydroperoxide 447 Testicular hyaluronidase 71 tetrodotoxin (TTX) 379, 474 Thyroid hormone receptor (TRβ2) 43, 46. 49. 262, 359 Toll-like receptor 4 78 Transcription factor Orthodenticle (Otd) 258 Transcription factor Rx 100 Transducin 8, 127, 144 Transient (Y) fashion 299 Transient receptor potential (TRP) 210

Index Transient receptor potential canonical (TRPC) 200 Trichromacy 308 Trichromatic color theory 310 Tritan defects 318 Tritanopia 318 TRP channel 271, 276 TRP protein family 269 TRP/TRPL channels 251 TRP-INAD interaction 274, 276 Tyrosine phosphatase SHP 133 U Ultraviolet-visible (UV/Vis) spectrophotometry 180 Univariance 308, 333 UV and M opsins 358 UV-sensitive cones 355 V V*(λ) 336 V10 (λ) 336 Vascular endothelial growth factor (VEGF) 409, 438 Vasoactive intestinal polypeptide (VIP) 484 Vasopressin V2R receptor 183 VAX 22 Vax2 21 Ventral lateral geniculate (VLG) 471 Verapamil 212 VERIS 375 Vigabatrin 386 Visual acuity 332 Visual cycle 69, 88 Visual effectiveness 331 Visual field 243 Visual G protein transducin 275, 269 Visual perception 226 Vitamin A 45, 91 Vitiligo mice 72 Vitreous 5 Voltage-gated Ca2+ channels (VGCCs) 202 Vsx1 41 W Wide-field amacrine cells 293 Wnt-inhibitory factor (WIF-1) 46 Wnts 46

Index X X-ray crystallography 179 Y y ommatidia 253 Z Zeaxanthin 18, 397 ZO-1 271 α-helix 232

509 α-transducin 307 αvβ5 integrin 72, 79 β5 integrin 73, 81 β-adrenergic receptor 181 b-catenin 46 β-oleoyl-γ-palmitoyl-L-αphosphatidylcholine 443 γ-aminobutyric acid (GABA) 47, 293, 469, 484 ω-conotoxin 205